Download Persistence of Disease Agents in Carcases / Animal

Persistence of Disease
Agents in Carcases and
Animal Products
Report for Animal Health Australia
by
Scott Williams Consulting Pty Ltd
Revised - December 2003
Published DECEMBER 2003
© Animal Health Australia 2003
ISBN 1 876714 59 X
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Table of Contents
Introduction....................................................................................................................... 5
Purpose of Consultancy .............................................................................................. 5
Methodology ............................................................................................................... 6
Notes on the Report .................................................................................................... 7
Review of Disease Agents ................................................................................................. 8
Bunyaviridae ............................................................................................................... 8
Nairobi sheep disease.............................................................................................. 8
Rift Valley fever ................................................................................................... 10
Flaviviridae ............................................................................................................... 13
Japanese encephalitis ............................................................................................ 13
Tickborne encephalitides ...................................................................................... 15
Wesselsbron disease.............................................................................................. 18
Orthomyxiviridae...................................................................................................... 20
Avian influenza (highly pathogenic) .................................................................... 20
Equine influenza ................................................................................................... 24
Swine influenza..................................................................................................... 26
Paramyxoviridae ....................................................................................................... 28
Hendra virus.......................................................................................................... 28
Menangle virus (porcine paramyxovirus)............................................................. 29
Newcastle disease ................................................................................................. 30
Nipah virus............................................................................................................ 34
Peste des petits ruminants ..................................................................................... 35
Rinderpest ............................................................................................................. 38
Picornaviridae ........................................................................................................... 42
Foot-and-mouth disease ........................................................................................ 42
Swine vesicular disease......................................................................................... 51
Teschen disease (porcine polioencephalomyelitis)............................................... 54
Poxviridae ................................................................................................................. 56
Lumpy skin disease............................................................................................... 56
Sheep pox.............................................................................................................. 59
Reoviridae ................................................................................................................. 63
African horse sickness .......................................................................................... 63
Bluetongue ............................................................................................................ 65
Equine encephalosis.............................................................................................. 68
Retroviridae............................................................................................................... 69
Jembrana disease................................................................................................... 69
Maedi-visna........................................................................................................... 71
Pulmonary adenomatosis ...................................................................................... 74
Rhabdoviridae ........................................................................................................... 76
Australian lyssavirus............................................................................................. 76
Rabies.................................................................................................................... 77
Vesicular stomatitis............................................................................................... 80
Togaviridae ............................................................................................................... 83
Getah virus disease ............................................................................................... 83
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Porcine reproductive and respiratory syndrome ................................................... 84
Western, Eastern and Venezuelan equine encephalomyelitis............................... 88
Other viruses ............................................................................................................. 89
African swine fever............................................................................................... 89
Aujeszky’s disease ................................................................................................ 92
Borna disease ........................................................................................................ 96
Classical swine fever............................................................................................. 98
Infectious bursal disease (hypervirulent form) ................................................... 102
Transmissible gastroenteritis .............................................................................. 105
Vesicular exanthema........................................................................................... 108
Prions ...................................................................................................................... 111
Bovine spongiform encephalopathy and scrapie ................................................ 111
Bacteria, mycoplasmas and fungi ........................................................................... 116
Anthrax ............................................................................................................... 116
Heartwater........................................................................................................... 120
Brucellosis (B. abortus) ...................................................................................... 122
Brucellosis (B. melitensis) .................................................................................. 126
Bovine tuberculosis............................................................................................. 130
Haemorrhagic septicaemia.................................................................................. 136
Glanders .............................................................................................................. 138
Contagious equine metritis ................................................................................. 140
Potomac fever ..................................................................................................... 142
Contagious bovine pleuropneumonia ................................................................. 143
Epizootic lymphangitis ....................................................................................... 145
Protozoa .................................................................................................................. 147
Surra.................................................................................................................... 147
Dourine ............................................................................................................... 150
East Coast fever .................................................................................................. 152
Equine babesiosis (piroplasmosis) ...................................................................... 154
Multicellular parasites............................................................................................. 155
Sheep scab........................................................................................................... 155
Tropilaelaps mite ................................................................................................ 157
Varroa mites........................................................................................................ 158
Acarapis (tracheal) mite...................................................................................... 160
Screw-worm fly .................................................................................................. 161
Braula fly ............................................................................................................ 163
Small hive beetle................................................................................................. 164
Trichinellosis....................................................................................................... 166
Appendix 1..................................................................................................................... 170
Example search strategy ......................................................................................... 170
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Introduction
Purpose of Consultancy
The purpose of this consultancy was to provide information to enable Commonwealth and
State/Territory veterinary, environmental and public health officials to review animal disposal
methods in Australia and to augment the Disposal manual of AUSVETPLAN.
Terms of reference for the consultancy were as follows:
For each of the 63 disease agents identified in the Cost Sharing Agreement, their
persistence in carcasses and animal products, both after slaughter and when the carcases
and animal products are disposed of or treated by the methods identified in the scoping
workshop of 2-3 July 2001*, is to be determined.
A listing of all 63 disease agents is to be constructed in a user-friendly, but scientifically
accurate, format detailing the minimum requirements for inactivation / destruction of the
disease agent, designed to be used for the purpose of determining the most appropriate
method of disposal of infected carcasses and animal products.
* These methods are:
•
Left in situ, i.e. no further treatment
•
Burial
•
Above-ground ‘burial’
•
Aerobic decomposition
•
Burial at sea
•
Chilling
•
Rendering
•
Combustion.
The Commonwealth/States/Territories gives no warranty that the information contained in AUSVETPLAN
is correct or complete. The Commonwealth shall not be liable for any loss howsoever caused whether due
to negligence or other arising from use or reliance on this code.
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Methodology
The consultancy was undertaken in four overlapping stages:
1. Review of microbiological and other texts for information relating to persistence
and disposal methods for the 63 organisms.
Microbiological texts were generally found to be unrewarding. By far the most useful text for
most of the agents studied in this review was Infectious diseases of livestock with special
reference to Southern Africa, vols I and II, eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C.,
Oxford University Press, Cape Town, 1994.
Other texts commonly referred to were Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic
diseases of animals: a field guide for Australian veterinarians, Australian Government Publishing
Service, Canberra, and Radostits, O.M., Gay, C.G., Blood, D.C., & Hinchcliff, K.W. (eds) 2000,
Veterinary medicine, ninth edition, W. B. Saunders Company Ltd, London.
2. Detailed literature search.
A search was made of Medline (PubMed), CAB Abstracts, Agricola, Australian Bibliography of
Agriculture (ABOA) and Current Contents databases for relevant papers (reviews,
epidemiological articles) on each disease / agent. The searches were conducted by the Senior
Librarian at the Victorian Institute of Animal Science. The search strategy was similar for each
disease / agent, although more focused in the case of diseases such as foot-and-mouth disease.
An example of the search strategy is given in Appendix 1.
Key reviews and recent references were acquired and checked for information not obtained in
stage 1. It was outside the scope of the consultancy to obtain every reference identified as
relevant. As far as possible throughout this report, original references are identified even where
they have been cited from other papers. The source of the citation is also given.
During the review, extensive cross-checking was made against draft and final Import Risk
Assessments (IRAs) prepared over the last two years by the Australian Quarantine and Inspection
Service (AQIS) / Animal Biosecurity. The IRAs provide a very thorough review of the
persistence and inactivation of a range of disease agents in selected products, and analysis of the
risk of transmission by those products.
3. Telephone and/or e-mail contact with individual experts and other key persons.
Australian and international experts for most diseases were contacted directly. The experts were
asked for suggestions on key references, to cross-check the literature reviews; and about
information from unpublished or in-progress studies. Names were initially obtained from the
Office International des Epizooties (OIE) web site. Others were identified from papers obtained
through stages 1 and 2, or from referrals.
Contacting experts proved to be much more time-consuming than anticipated. Some experts were
very difficult to contact, despite repeated telephone calls and e-mails, and a number were not
reached within the timeframe of the consultancy.
4. Compilation of report.
This report was first delivered in December 2001. This update, with modification to the
conclusions, was completd in December 2003.
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Notes on the Report
1. This is a technical report focusing on the persistence and inactivation of disease agents.
Greater emphasis has been placed on thorough review of the relevant literature than on
drawing conclusions about disposal methods. A number of factors will contribute to
determining the optimum disposal method, including effectiveness in halting transmission,
cost, attitude to risk, ease of implementation, public perception, and local factors (such as
topography, accessibility, and native and feral fauna). These factors were outside the scope
of this review.
2. There are many gaps in the knowledge about persistence and inactivation of certain disease
agents in carcases and animal products. In particular, there very few primary studies on the
inactivation of agents in disposal methods other than leaving in situ, incineration and burial.
A number of the experts contacted commented on the paucity of information available.
Reasons for this lack of information include low relevance to the epidemiology or control of a
number of the diseases examined. For example, for most of the vector-borne diseases, the
vectors have appropriately received most of the attention. Information on persistence may
also be a low priority to countries in which the disease is endemic, and where efforts are
therefore directed at control (e.g. vaccines) rather than stamping out. At the same time, there
is reluctance to work with the agent in countries where it is exotic.
3. For each of the disease agents in the report, the following information is presented:
• General characteristics of the agent and of the disease that are relevant to transmission via
carcases and animal products – including in vitro data on physical and chemical
characteristics, survival in the environment, effective disinfectants, where the agent is
found in the body, and how it is transmitted
• Summaries of specific studies on the persistence and inactivation of the agent in products
from the host, with some analysis and interpretation, including information from experts
where applicable
• Conclusions based on the available data for disposal of carcases and products
• References and expert contacts made.
4. This revised version of the report departs from the letter of the original terms of reference.
The paucity of literature, for virtually all of the agents studied, precluded firm conclusions
being drawn about persistence following disposal by each of the methods presented. The
‘Conclusions’ section for each agent has been revised, so that it now summarises key points
and makes recommendations based only on known information and reasonable
extrapolation.These recommendations should not necessarily be considered as policy
statements by Animal Health Australia, the Australian or State/Territory governments.
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Review of Disease Agents
Bunyaviridae
Nairobi sheep disease
Agent:
Family Bunyaviridae, genus nairovirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: Nairobi sheep disease virus is a relatively unstable agent. At its optimum
pH range of 7.4-8.0 and with 2% serum, it has a half-life of 6.8 days at 0oC and 1.5 hours at 37oC.
Stability is greater in citrated whole blood, where the presence of protein is protective: 66 days at
5oC, 33 days at 15-18oC, 42 hours at 37oC, 1.5 hours at 50oC, and 5 minutes at 60oC. NSDV is
sensitive to lipid solvents and detergents and is rapidly inactivated outside the optimum pH range
(Terpstra 1990, Montgomery 1917).
NSD is transmitted by ticks, mainly of the genus Rhipicephalus (Terpstra 1990).
Carcases and meat products: The pathogenesis of NSD includes a viraemic phase in which the
virus spreads to most internal organs. No references were uncovered on persistence of NSDV in
carcases or muscle during the conduct of this review. The virus would be expected to disappear
quickly from the carcase as the pH dropped with rigor mortis. AQIS (1999a) does not consider
NSD to be a risk in imported meat products.
Davies (1988) noted that, in contrast to Rift Valley fever, there were no reports of humans
becoming infected despite handling infected carcases.
Milk and milk products: No reports were uncovered on the shedding of NSDV in milk. AQIS
(1999b) does not regard milk products as posing a threat for the importation of NSD.
Skins, hides and fibres: No reports were uncovered on the contamination of skins, wool or hair
by NSDV. In a draft assessment, AFFA (2001) has determined that skins and hides do not pose
an import risk for NSD.
Semen/embryos: No reports were uncovered on the shedding of NSDV in semen or with
embryos.
Faeces: Stock affected with NSD suffer an acute diarrhoea (Terpstra 1990), but there are no
reports of virus being isolated from faeces.
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Conclusions:
•
Nairobi sheep disease virus is a fragile virus outside its narrow optimal pH range. It
is inactivated by the pH changes associated with rigor mortis in meat.
•
NSDV is spread by a tick vector.
•
Carcases and animal products present a negligible disease risk and no special
precautions for disposal are warranted.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999a, Importation of sausage casings into
Australia, import risk analysis, December 1999, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999b, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
Davies, F.G. 1988, ‘Nairobi sheep disease’, in The arboviruses: epidemiology and ecology. Vol
III, ed. Monath, T.P., CRC Press, Boca Raton, Florida, pp. 191-203.
Montgomery, R.E. 1917, ‘On a tick-borne gastro-enteritis of sheep and goats occurring in British
East Africa’, J Comp Pathol Ther, vol. 30, pp. 28-58, cited in Terpstra 1990.
Terpstra, C. 1990, ‘Nairobi sheep disease virus’, in Virus infections of ruminants, eds. Z. Dinter
and B. Morein, Elsevier Science Publishers B.V., Amsterdam, pp. 495-500.
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Rift Valley fever
Agent:
Family Bunyaviridae, genus phlebovirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: Rift Valley fever virus is very stable in liquid media (blood or serum)
under various conditions of refrigeration, lasting several months at 4oC and three hours at 56oC.
It is stable in aerosol at 23oC and 50-85% humidity. RVFV is stable between pH 7-9 but rapidly
inactivated below pH 6.8. It is also inactivated by lipid solvents, including ether and sodium
deoxycholate, and low concentrations of formalin (Swanepoel and Coetzer 1994).
RVF is transmitted by a number of mosquitoes, most notably culicines (Geering et al 1995).
Carcases and meat products: There are many reports of humans, such as abattoir workers,
becoming infected with RVF after contact with infected tissues. No outbreaks have occurred in
urban consumer populations. It is generally considered that meat from RVF-infected animals is
not a source of transmission, as the pH changes associated with rigor mortis inactivate the virus
(Chambers and Swanepoel 1980, MacDiarmid and Thompson 1997, Swanepoel 1981). The OIE
were recently advised of this position (Gerdes, pers comm).
AQIS (1999a) does not consider RVF to be a risk in imported meat products. MacDiarmid and
Thompson (1997) concluded that sheep and goat meat presented little risk for the international
spread of RVF.
Milk and milk products: RVFV has been detected in low concentration in milk. Infected milk
may have been connected with an outbreak of RVF in humans in Mauritania (Swanepoel and
Coetzer 1994). DPIE (1996) states that pasteurisation or treatment with acid inactivates the virus.
AQIS (1999b) does not regard milk products as posing a threat for the importation of RVF.
Skins, hides and fibres: No specific references were found on this aspect of the disease. In a draft
assessment, AFFA (2001) has determined that skins and hides do not pose an import risk for
RVF. AUSVETPLAN (DPIE 1996) notes that little is known about the persistence of RVFV in
these products but that there is some small risk. It advocates the burial or disinfection of skins
and scouring (+/- carbonisation) of fibres.
Semen/embryos: No specific references were found on this aspect of the disease. Radostits et al
(2000) state that “other than milk and aborted foetuses no body secretions or excretions contain
the virus”. However, DPIE (1996) notes that RVFV is likely to be present in semen and that
transmission is possible by this means, and that the virus is known to be present on ova but is
probably not transmitted.
Faeces: Similar comments apply to those regarding skins, hides and fibres. No specific
references were found.
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Conclusions:
•
Rift Valley fever virus can readily survive in body fluids between pH7-9, but is
fragile outside this pH range. It is inactivated by the pH changes associated with
rigor mortis in meat.
•
RVF is principally spread by mosquitoes, but human infection can also occur by
contacting fresh tissues or body fluids from an infected animal.
•
Burial, burning and rendering are the preferred methods for disposal of fresh
carcases.
•
Meat, skins, hides and fibres present a negligible transmission risk.
•
Milk should be pasteurised to inactivate the RVF virus.
•
Semen and embryos should be destroyed.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999a, Importation of sausage casings into
Australia, import risk analysis, December 1999, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999b, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
Chambers, P.G., & Swanepoel, R. 1980, ‘Rift Valley fever in abattoir workers’, Cent Afr J Med,
vol. 26, pp. 122-126, cited in Swanepoel and Coetzer 1994.
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Rift Valley Fever, DPIE, Canberra.
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
MacDiarmid, S.C., & Thompson, E.J. 1997, ‘The potential risks to animal health from imported
sheep and goat meat’, Rev sci tech Off int Epiz, vol. 16(1), pp.45-56.
Radostits, O.M., Gay, C.G., Blood, D.C., & Hinchcliff, K.W. (eds) 2000, Veterinary medicine,
ninth edition, W. B. Saunders Company Ltd, London.
Swanepoel, R. 1981, ‘Observations on Rift Valley fever in Zimbabwe’, Contributions to
Epidemiology and Biostatistics, vol. 3, pp. 83-91, cited in Swanepoel and Coetzer 1994.
Swanepoel, R., & Coetzer, J.A.W. 1994, ‘Rift Valley fever’, in Infectious diseases of livestock,
Vol I, eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape
Town, pp. 688-717.
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Experts contacted:
Dr Truuske Gerdes
Onderstepoort Veterinary Institute
Private bag X05
Onderstepoort 0110
SOUTH AFRICA
P +27 12 529 9114
F +27 12 529 9418
[email protected]
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Flaviviridae
Japanese encephalitis
Agent:
Family Flaviviridae, genus flavivirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: Japanese encephalitis virus is extremely fragile, and does not survive off
the host for more than a few hours (DPIE 1998).
Transmission of JEV is principally by Culex spp mosquitoes. Pigs are amplifiers of the virus and
the only domestic livestock species to develop viraemia of sufficient titre to infect mosquitoes.
Virus can be isolated from CNS tissue, CSF and serum (Ellis et al 2000).
Carcases and meat products: No references to this aspect of the disease were found in the
literature. AUSVETPLAN (DPIE 1998) stipulates no special precautions for the disposal of
carcases from infected premises.
Milk and milk products: Inapparent infections may occur in species producing milk as a product
for human consumption (cattle, sheep, goats) (DPIE 1998). No reports were found in the
literature regarding the presence or persistence of JEV in milk.
Skins, hides and fibres: In a draft assessment, AFFA (2001) has concluded that the likelihood of
JEV transmission via skins, hides or fibres would be negligible.
Semen/embryos: JE has been experimentally transmitted to gilts in boar semen (Habu et al 1977).
AQIS (2000) has determined that pig semen poses a moderate quarantine risk for JEV.
Faeces: No specific reports were found of the presence or persistence of JEV in faeces.
Conclusions:
•
Japanese encephalitis virus is a fragile virus which can only survive for a few hours
away from an animal host.
•
JEV is spread by a mosquito vector.
•
Carcases and animal products present a negligible transmission risk and no special
precautions for disposal are warranted.
•
Semen and milk from infected or suspect animals should not be used.
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References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 2000, Draft porcine semen risk analysis:
risk assessment and risk management options, March 2000, AQIS, Canberra.
DPIE (Department of Primary Industries and Energy) 1998, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Japanese encephalitis, DPIE, Canberra.
Ellis, P.M., Daniels, P.W., & Banks, D.J. 2000, ‘Japanese encephalitis’, Vet Clin Nth Am Eq
Pract, vol. 16, pp. 565-578.
Habu, A., Murakami, Y., Ogasa, A., & Fujisaka, Y. 1997, ‘Disorder of spermatogenesis and viral
discharge into semen in boars affected with Japanese encephalitis virus’, Virus (Tokyo), vol. 27,
pp. 21-26, cited in AQIS 2000.
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Tickborne encephalitides
Agent:
Family Flaviviridae, genus flavivirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: Tickborne encephalitis complex viruses are not very resistant to
environmental influences. Tickborne flaviviruses are reported to be sensitive to pH below 6.0
(compared to mosquito-borne flaviviruses, which are sensitive below pH 8.0). They are
inactivated by heating to 56oC for 30 minutes but are stable at temperatures below -60oC.
TBE complex viruses are inactivated by UV and gamma radiation and by proteases, lipid solvents
and detergents, as well as by low concentrations of aldehydes, halogens, hydrogen peroxide, and
beta-propiolactone (Gresikova and Kaluzova 1997, Swanepoel 1994, Reid pers comm, Gould
pers comm).
Carcases and meat products: No published reports were found on this aspect of the disease.
Gould (pers comm) believes that louping-ill virus would lose its infectivity rapidly (within a day
or two) due to putrefaction and bacteria. Inactivation would be slowed by low temperatures.
Some virus could be washed out of the carcase but it is likely to be quickly inactivated (days, or
probably hours) and diluted below risk level. Similar advice was offered by Reid (pers comm).
Ixodes ticks are normally responsible for transmission. However, there is a report in which 10 out
of 16 piglets appear to have become infected through ingestion of the virus (Bannatyne et al
1980). Two to three weeks previously, the piglets had been fed the uncooked carcases of lambs
that had died with signs of louping-ill. Grouse chicks have also been infected with louping-ill
virus after ingesting infected ticks (Gould pers comm). Although a minor aspect of the
epidemiology, these findings would suggest the preferential burial of carcases rather than leaving
in situ.
Milk and milk products: Transmission of louping-ill virus from goat dam to kid via milk has been
demonstrated by Reid et al (1984). The finding suggested a public health risk consistent with
known European cases of human infection by other members of the TBE complex from drinking
goat’s milk. Outbreaks of disease have also been linked to the milk of infected sheep (Gresikova
et al 1975) and cattle (Jezyna 1976). The virus is totally inactivated by pasteurisation using
“normal” pasteurisation temperature / time treatments (Reid pers comm).
Skins, hides and fibres: The risk of infective TBE complex virus on skins, hides or fibres is
negligible, given its relative fragility and normal mode of transmission via ticks (AFFA 2001).
Semen/embryos: No reports of TBE complex virus in semen or embryos were found during this
review, although Papadopoulos et al (1971) reported the isolation of a TBE complex virus from a
flock of goats with abortions. AQIS (2000) concluded that “the risk estimate for [louping-ill and
related viruses] without risk management measures is negligible for semen and embryos”.
Faeces: No references were found on the presence or persistence of TBE complex virus in faeces
during this review.
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Conclusions:
•
Tickborne encephalitis complex viruses are sensitive agents which are unlikely to
survive for more than a few days in animal products in the field.
•
TBEC viruses are normally spread by a tick vector, but transmission via ingestion
cannot be ruled out.
•
Carcases and animal products present a low transmission risk but should be
disposed of using a method that prevents scavenging during the first week.
•
Milk should only be used following pasteurisation.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 2000, An analysis of the disease risks, other
than scrapie, associated with the importation of ovine and caprine semen and embryos from
Canada, the United States of America and member states of the European Union, final report,
AQIS, Canberra.
Bannatyne, C.C, Wilson, R.L., Reid, H.W., Buxton, D., & Pow, I. 1980, ‘Louping-ill virus
infection of pigs’, Vet Rec, vol. 106, p. 13.
Gresikova, M., & Kaluzova, M. 1997, ‘Biology of tick-borne encephalitis virus’, Acta virologica,
vol. 41, pp. 115-124.
Gresikova, M., Sekeyova, M., Stupalovaa, S., & Necas, S. 1975, ‘Sheep milk borne epidemic of
tick-borne encephalitis in Slovakia’, Intervirology, vol. 5, p. 57, cited in Gresikova and Kaluzova
1997.
Jezyna, C. 1976, ‘A milk-borne outbreak of tick-borne encephalitis in the Olsztyn province’,
Przegl Epidemiol, vol. 30, p. 479, cited in Gresikova and Kaluzova 1997.
Papadopoulos, O., Paschaleri-Papadopoulos, E., Deligaris, N., & Doukas, G. 1971, ‘[Isolation of
tick-borne encephalitis virus from a flock of goats with abortions and fatal disease (a preliminary
report)]’, Kteniatrika-Nea, vol. 3, pp. 112-113, cited in AQIS 2000.
Reid, H.W., Buxton, D., Pow, I., & Finlayson, J. 1984, ‘Transmission of louping-ill virus in goat
milk’, Vet Rec, vol. 114, pp. 163-165.
Swanepoel, R. 1994, ‘Louping ill’, in Infectious diseases of livestock, Vol II, eds. Coetzer,
J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 671-677.
Persistence of Disease Agents
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Experts contacted:
Dr Earnest Gould
Centre for Ecology and Hydrology
Mansfield Rd
Oxford OX1 3SR Oxon
UK
P +44 8165 281630
F +44 8165 281696
[email protected]
Dr Hugh Reid
Moredun Research Institute
Pentlands Science Park
Bush Loan
Midlothian EH26 0PZ
Scotland
UK
P +44 131 445 5111
F +44 131 445 6111
[email protected]
Persistence of Disease Agents
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Wesselsbron disease
Agent:
Family Flaviviridae, genus flavivirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: According to Swanepoel and Coetzer (1994), Wesselsbron disease virus
has not been well characterised, but it has the properties typical of haemagglutinating flaviviruses.
These are sensitive to acidity (< pH 8.0), temperatures above 40oC, lipid solvents and detergents
(Gresikova and Kaluzova 1997, Reid pers comm, Gould pers comm).
Transmission of the disease is principally by Aedes spp mosquitoes. Handling of infected
carcases has caused infection in humans, and aerosol transmission may be possible, but it appears
there is no direct spread between sheep (Swanepoel and Coetzer 1994).
Carcases and meat products: No published reports were found on this aspect of the disease.
Humans have been reported to have contracted the disease after performing post mortems on
infected animals (Heymann et al 1958).
Milk and milk products: No reports were found on the detection or transmission of WDV in milk.
AQIS (1999) has listed Wesselsbron disease virus as an agent that may be excreted in milk, but
which is not considered to pose a quarantine hazard in milk or milk products.
Skins, hides and fibres: The risk of infective WDV on skins, hides or fibres is negligible, given its
relative fragility and normal mode of transmission (AFFA 2001).
Semen/embryos: No reports were found on the detection or transmission of WDV in semen or
embryos.
Faeces: No reports were found on the detection or transmission of WDV in faeces.
Conclusions:
•
Wesselsbron disease virus is a sensitive agent which is unlikely to survive for more
than a few days in carcases or animal products in the field.
•
Wesselsbron disease is principally spread by mosquitoes, but human infection can
also occur by contacting fresh tissues or body fluids from an infected animal.
•
Carcases and animal products present a low transmission risk but should be
disposed of using a method that prevents scavenging.
Persistence of Disease Agents
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References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
Gresikova, M., & Kaluzova, M. 1997, ‘Biology of tick-borne encephalitis virus’, Acta virologica,
vol. 41, pp. 115-124.
Heymann, C.S., Kokernot, R.H., & De Meillon, B. 1958, ‘Wesselsbron virus infections in man’,
Sth Afr Med J, vol. 32, pp. 543-545, cited in Swanepoel and Coetzer 1994.
Swanepoel, R., & Coetzer, J.A.W. 1994, ‘Wesselsbron disease’, in Infectious diseases of
livestock, Vol II, eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press,
Cape Town, pp. 663-670.
Experts contacted:
Dr Earnest Gould
Centre for Ecology and Hydrology
Mansfield Rd
Oxford OX1 3SR Oxon
UK
P +44 8165 281630
F +44 8165 281696
[email protected]
Dr Hugh Reid
Moredun Research Institute
Pentlands Science Park
Bush Loan
Midlothian EH26 0PZ
Scotland
UK
P +44 131 445 5111
F +44 131 445 6111
[email protected]
Persistence of Disease Agents
Revised December 2003
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Scott Williams Consulting Pty Ltd
Orthomyxiviridae
Avian influenza (highly pathogenic)
Agent:
Family Orthomyxiviridae, type A
Agent type:
Virus
Persistence and inactivation:
Highly pathogenic avian influenza virus survives well in organic matter and in water. It has been
isolated from water contaminated with faeces for up to four days at 22oC, and for more than 30
days at 0oC (Webster et al 1978), while Stallknecht et al (1990) estimated retention of infectivity
for up to 207 days at 17oC and 102 days at 28oC in lake water. HPAIV is stable between pH 5.5
and 8 (DPIE 1996). The wide variation in heat lability between strains of the virus has been
noted by Blaha (1989), who reported survival of between 15 minutes and six hours at 56oC.
Storage of the virus at -70oC or lyophilisation is recommended for long-term maintenance of
infectivity (Easterday et al 1997).
HPAIV is destroyed by lipid solvents such as detergents, as well as formalin, beta-propiolactone,
oxidising agents, dilute acids, ether, sodium desoxycholate, hydroxylamine, sodium
dodecylsulphate, and ammonium ions (Easterday et al 1997).
HPAI is highly contagious. It is excreted in high concentrations in faeces, and in oral and nasal
discharges. Spread is by direct contact, by movement of infected birds, and by fomites (Geering
et al 1995). The virus is found in the brain, skin, and most visceral organs (Swayne and Suarez
2000).
Carcases and meat products: Swayne and Suarez (2000) note, with regard to the risk of importing
HPAI in meat products, that “HPAI is a systemic disease and the virus can be present in most
tissues, including meat”. AFFA (2001) has determined that there is sufficient prima facie
evidence to justify a full risk assessment on HPAI in uncooked chicken meat. The report cites
Becker and Uys (1967), who isolated the virus up to six days post-inoculation from the muscle of
experimentally infected chickens.
DPIE (1996) states that the virus survives several days in carcases at ambient temperatures and up
to 23 days when refrigerated. It cites the following minimum core temperatures to kill HPAIV:
• 70oC for 30 minutes
• 75oC for 5 minutes
• 80oC for 1 minute.
Senne et al (1994) found evidence that composting may be effective in inactivating HPAIV in
carcases. The composting was performed in bins at an ambient temperature of 22oC. Various
organs of experimentally-infected birds were composted with poultry carcases in goat manure and
straw in a volume ratio of 1:2:1. The samples were composted in two stages of ten days each,
with a turning of the pile at 10 days.
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The upper layer of the compost reached 57.3oC during the first stage and 41.5oC during the
second. Corresponding peak temperatures for the lower layer were 58.3oC and 42.8oC. HPAIV
was not isolated at the end of ten days or at twenty days.
Eggs and egg products: Vertical transmission of HPAI through embryonated eggs has not been
demonstrated (Geering et al 1995). However, the virus has been detected in and on eggs from
naturally infected chickens, arising from deposition during egg formation, and as an external
contaminant (Capucci et al 1985). No references were found on the persistence of the virus in
eggs.
A number of approaches effectively inactivate AIV in eggs (King 1991). Ackland et al (1985)
heated egg waste from influenza vaccine production at 60oC for three hours. King (1991)
examined stability of HPAIV at pasteurisation temperatures. Virus was diluted 10-2 in egg yolk,
albumen, or allantoic fluid. HPAIV was inactivated in yolk at 57oC and in allantoic fluid at 62oC
in less than 5 minutes. Virus in albumen was inactivated in 5-10 minutes at 57oC.
The author noted current (1991) US recommended high-temperature, short-time pasteurisation
times: 3.5 minutes at 57oC for albumen, at 60oC for whole egg, and at 61oC for yolk respectively,
or whole eggs at 56oC for 35 minutes, 57oC for 15 minutes, or 60oC for 3.5 minutes. He
concluded that these combinations would be marginal even for HPAIV (Newcastle disease virus
was more hardy) and supported recommendations for their increase. DPIE (1996) states that 4.5
minutes at 64oC is thought to kill HPAIV but that 2.5 minutes is insufficient.
In another experiment, heat and chemical effects on allantoic fluid containing virus, with or
without chicken serum, were evaluated. Infectivity of HPAIV was eliminated after 60 minutes at
56oC, and after 30 minutes at 60oC. Formalin 0.01% destroyed HPAIV in serum. Betapropiolactone 0.1% destroyed the virus in both media, and binary ethyleneimine 0.01M was
effective after 5 hours (King 1991).
Other products: Virus can be present in faeces in concentrations as high as 107 infectious particles
per gram, and may persist for more than 44 days (Utterback 1984). HPAIV has been recovered
from liquid manure 105 days after depopulation. In other reports, the virus retained infectivity in
faeces for 35 days at 4oC and for 7 days at 20oC. The presence of organic material protects the
virus. Manure can be disposed of by burial, composting in a pile covered by plastic, or
rototilling1 (Easterday et al 1997).
Conclusions:
1
•
Highly pathogenic avian influenza virus can survive in organic matter and water for
months. However, the virus is quite sensitive to heat and is inactivated by a wide
range of disinfectants.
•
The HPAI virus is highly contagious. Biosecurity of potentially infective material is
most important.
•
Burning and burial are the preferred methods for disposal of infective material.
A definition for this word could not be found
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References:
Ackland, N.R., Tannock, G.A., & Young, I.F., 1985, ‘A device for the nondestructive
decontamination of large volumes of infected egg waste’, Appl Environ Microbiol, vol. 49, pp.
920-924, cited in King (1991).
Alexander, D.J. 1995, ‘The epidemiology and control of avian influenza and Newcastle disease’,
J Comp Path, vol. 112, pp. 105-126.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2000, The importation of
non-viable eggs and products containing egg. Technical issues paper, December 2000, AFFA,
Canberra.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Generic import risk
analysis (IRA) for uncooked chicken meat. Issues paper, July 2001, AFFA, Canberra.
Becker, W.B., & Uys, C.J. 1967, ‘Experimental infection of chicken with influenza A/Tern/South
Africa/1961 and chicken/Scotland/1959 viruses. 1.Clinical picture and virology’, J Comp Pathol,
vol. 77, pp. 159-165, cited in AFFA 2001.
Blaha T., 1989, ‘Fowl plague’, in Applied veterinary epidemiology, Elsevier, Amsterdam, pp. 6973, cited in AFFA 2000.
Cappucci, D.T. Jr., Johnson, D.C., Brugh, M., Smith, T.M., Jackson, C.F., Pearson, J.E., &
Senne, D.A. 1985, ‘Isolation of avian influenza virus (subtype H5N2) from chicken eggs during a
natural outbreak’, Avian Diseases, vol. 29, pp. 1195-1200, cited in King 1991 and AFFA 2000.
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Avian Influenza, DPIE, Canberra.
Capucci, Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field
guide for Australian veterinarians, Australian Government Publishing Service, Canberra.
Easterday, B.C., Hinshaw, V.S., & Halvorson, D.A. 1997, ‘Influenza’, in Diseases of poultry,
tenth edition, ed. Calnek, B.W., Mosby-Wolfe, London, pp. 583-605.
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
King, D.J. 1991, ‘Evaluation of different methods of inactivation of Newcastle disease virus and
avian influenza virus in egg fluids and serum’, Avian Dis, vol. 35, pp. 505-514.
Senne, D.A., Panigrahy, B., & Morgan, R.L. 1994, ‘Effect of composting poultry carcasses on
survival of exotic avian viruses: highly pathogenic avian influenza (HPAI) virus and adenovirus
of egg drop syndrome-76’, Avian Dis, vol. 38, pp. 733-737.
Stallknecht, D.E., Shane, S.M., Kearney, M.T., & Zwank, P.J. 1990, ‘Persistence of avian
influenza viruses in water’, Vet Res Comm, vol. 12, pp. 125-141, cited in Alexander 1995.
Swayne, D.E., & Suarez, D.L. 2000, ‘Highly pathogenic avian influenza’, Rev sci tech Off int
Epiz, vol. 19(2), pp. 463-482.
Persistence of Disease Agents
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Utterback, W. 1984, ‘Update on avian influenza through February 21, 1984 in Pennsylvania and
Virginia’, in Proc 33rd Western Poultry Dis Conf, pp. 4-7, cited in Alexander 1995.
Webster, R.G., Yakhno, M., Hinshaw, V.S., Bean, W.J., & Murti, K.G. 1978, ‘Intestinal
influenza: replication and characterisation of influenza viruses in ducks’, Virol, vol. 84, pp. 268276.
Experts contacted:
Dr Brundaban Panigrahy
National Veterinary Services Laboratories
Veterinary Services
Animal and Plant Health and Inspection Service
US Department of Agriculture
PO Box 844
Ames, Iowa 50010
USA
[email protected]
Persistence of Disease Agents
Revised December 2003
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Scott Williams Consulting Pty Ltd
Equine influenza
Agent:
Family Orthomyxiviridae, type A
Agent type:
Virus
Persistence and inactivation:
General characteristics: Equine influenza virus is enveloped and therefore does not survive long
outside the host (Mumford 1994). DPIE (1996) quotes persistence of 8-36 hours in the
environment.
The properties of equine influenza virus have been described by Yadav et al (1993). The virus
studied in that report was inactivated by exposure to UV light for 30 minutes or by heating at
50oC for 30 minutes. It was quickly inactivated by savlon, dettol, phenyl, alcohol, formalin, and
potassium permanganate but not by sodium carbonate (AUSVETPLAN (DPIE 1996) advocates
4% lysol). EIV persisted for up to 14 days in tap water at 4oC, or 2 days at 37oC, and in canal
water (where colloids were thought to be protective) for 1-2 weeks at 22oC or 37oC. It survived
in urine at pH 8.0 for up to 5 days and in soil at 18oC for one day.
EIV is spread via the respiratory route, almost exclusively by direct contact between horses.
Indirect spread by people and fomites may play some role (Mumford 1994).
Carcases and meat products: No reports on the persistence of equine influenza in horse carcases
were uncovered during the course of this review.
Skins, hides and fibres: EIV from infected aerosols might be expected to superficially
contaminate skins, but the unstable nature of the virus in the presence of UV light and heat means
that persistence for any period is highly unlikely. This assessment is confirmed by AFFA (2001).
Semen/embryos: EIV is present in the semen and embryos of infected horses, although
transmission is unlikely to occur by this route (DPIE 1996).
Faeces: No reports of EIV in faeces were found during this review. Faeces might be expected to
be contaminated by aerosols from infected animals. An outbreak of EI in China in 1987 was also
distinguished by signs of enteritis in affected animals (Hannant and Mumford 1996).
Conclusions:
•
Equine influenza virus is a sensitive virus which is unlikely to survive for more than a
few days away from an animal host. It is readily inactivated by exposure to UV light.
•
EIV is spread by the respiratory route, almost exclusively with close contact
between horses.
•
EIV may be present in semen and embryos from infected horses, but transmission is
unlikely to occur by this route.
•
Carcases and animal products present a negligible transmission risk and no special
precautions in disposal are warranted.
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References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Equine Influenza, DPIE, Canberra.
Mumford, J.A. 1994, ‘Equine influenza’, in Infectious diseases of livestock, Vol II, eds. Coetzer,
J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 854-859.
Hannant, D., & Mumford, J.A. 1996, ‘Equine influenza’, in Virus infections of vertebrates. Vol 6
Virus infections of equines, ed. Studdert, M.J., Elsevier Science Publishers, Amsterdam, pp. 285293.
Yadav, M.P., Uppal, P.K., & Mumford, J.A. 1993, ‘Physico-chemical and biological
characterisation of A/Equi-2 virus isolated from 1987 equine influenza epidemic in India’, Int J
Anim Sci, vol. 8, pp. 93-98.
Experts contacted:
Dr Thomas Chambers
OIE International Reference Laboratory for Equine Influenza
108 Gluck Equine Research Center
University of Kentucky
Lexington, Kentucky 40546-0099
USA
P +1 859 257 3407
F +1 859 257 8542
[email protected]
Dr Janet Daly (on behalf of Dr Jenny Mumford)
Centre for Preventive Medicine
Animal Health Trust
Lanwades Park
Kentford
Newmarket
Suffolk CB8 7UU
UK
P +44 1638 750659
F +44 1638 750794
[email protected]
Persistence of Disease Agents
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Swine influenza
Agent:
Family Orthomyxiviridae, type A
Agent type:
Virus
Persistence and inactivation:
General characteristics: Swine influenza virus is very labile, being rapidly inactivated by all
detergents and disinfectants. SIV is transmitted between pigs via the respiratory route. There is
no viraemia associated with infection, the virus localising to respiratory and lymphoid tissues
(Bachmann 1989).
Carcases and meat products: No reports of the presence of virus in meat from natural infection
were uncovered by this review, or by AFFA (2001a). Brown (pers comm) confirmed the lack of
data. The only reference of relevance is by Romjin et al (1989), who reported a study using pigs
inoculated with the H1N1 strain of SIV. Virus was recovered from lung stored at -20oC for
between 21 and 28 days, and also from blood for at least 14 days (when testing ceased). The
report also mentions sporadic recovery of virus from intestine, muscle and faeces.
The authors also examined the survival of virus on artificially contaminated samples of pig meat,
from which they were able to isolate virus for between 8 and 15 days at -20oC. Loss of infectivity
was slightly slower at 4oC compared to -20oC.
Skins, hides and fibres: SIV from infected aerosols might be expected to superficially contaminate
skins, but the unstable nature of the virus in the presence of UV light and heat means that
persistence for any period is highly unlikely. This assessment is confirmed by AFFA (2001b).
Semen/embryos: This review did not uncover any reports of SIV in semen, confirming the finding
of AQIS (2000).
Faeces: No reports were found of SIV excretion in faeces. Thomson (1994) cites a study by
Kawaoka et al (1987), in which only a human strain of influenza virus could be isolated from the
faeces of pigs, with two strains of SIV showing no evidence of replication in the intestinal tract.
Bøtner (1990) found that time to inactivation for SIV in slurry varied between 9 weeks at 5oC to
one hour at 50oC (detection limit 0.7 log10 TCID50/50L, initial dose 105.8 TCID50/50L).
Conclusions:
•
Swine influenza virus is a sensitive virus which is unlikely to survive for more than a
few days away from an animal host. It is readily inactivated by heat and exposure
to UV light.
•
SIV is spread by the respiratory route, with close contact required between infected
and susceptible pigs.
•
Carcases and animal products present a negligible transmission risk. The only
special precaution is that the method disposal must preclude scavenging by feral
pigs.
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References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001a, Generic import risk
analysis (IRA) for uncooked pig meat. Issues paper, January 2001, AFFA, Canberra.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001b, Import risk
analysis. Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 2000, Draft porcine semen risk analysis:
risk assessment and risk management options, March 2000, AQIS, Canberra.
Bachmann, P.A, 1989, ‘Swine influenza virus’, in Virus infections of vertebrates. Vol 2 Virus
infections of porcines, ed. Pensaert, M.B., Elsevier Science Publishers, Amsterdam, pp. 193-207.
Bøtner, A. 1990, Modelstudier vederonde overlevelse af virus i gylle under traditionel opbevaring
og under udradning i biogasanlaeg. Research report, State Veterinaary Institute for Virus
Research, Lindholm, Denmark, cited in Haas et al 1995.
Haas, B., Ahl, R., Bohm, R., & Strauch, D. 1995, ‘Inactivation of viruses in liquid manure’, Rev
sci tech Off int Epiz, vol. 14(2), pp. 435-445.
Kawaoka, Y., Bordwell, E., & Webster, R.G. 1987, Intestinal replication of influenza A viruses in
two mammalian species, Arch Virol, vol. 93, pp. 303-308, cited in Thompson 1994.
Romijn, P.C., Swallow, C., & Edwards, S. 1989, ‘Survival of influenza virus in pig tissues after
slaughter’, Vet Rec, vol. 124(9), p. 224.
Thomson, G.R. 1994, ‘Swine influenza’, in Infectious diseases of livestock, Vol II, eds. Coetzer,
J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 860-863.
Experts contacted:
Dr Ian Brown
Veterinary Laboratories Agency - Weybridge
New Haw, Addlestone
Surrey KT15 3NB
UK
P +44 1932 357339
F +44 1932 357239
[email protected]
Persistence of Disease Agents
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Paramyxoviridae
Hendra virus
Agent:
Family Paramyxoviridae, genus megamyxovirus (proposed name)
Agent type:
Virus
Persistence and inactivation:
Very little information is available on this virus. In experiments involving fruit bats, horses and
cats, the virus appeared to be poorly transmissible (Williamson et al 1998). The authors
concluded that respiratory spread or excretion via faeces was unlikely, and that the most plausible
theory for transmission was oral ingestion of urine-contaminated surfaces, water or feed.
However, the virus did not appear to persist long in the environment.
CSIRO at AAHL have assumed Hendra virus to have similar properties to Newcastle disease
virus until proven otherwise (Daniels pers comm). Virus numbers were reduced by about 6 logs
by pure methanol, and by 3% lysol after 3 minutes at room temperature. Anecdotally, the virus
appears stable when left in culture at room temperature (Abraham, pers comm).
Conclusions:
•
There is insufficient information available to make detailed recommendations on the
disposal or disinfection of animal carcases or products exposed to Hendra virus.
References:
Williamson, M.M., Hooper, P.T., Selleck, P.W., Gleeson, L.J., Daniels, P.W., Westbury, H.A.,
&Murray, P.K. 1998, ‘Transmission studies of Hendra virus (equine morbillivirus) in fruit bats,
horses and cats’, Aust Vet J, vol. 76, pp. 813-818.
Experts contacted:
Dr Peter Daniels and Mr Gordon Abraham
CSIRO Livestock Industries
Australian Animal Health Laboratory
Private Bag 24
5 Portarlington Rd
Geelong Vic 3220
P 03 5227 5000
F 03 5227 5555
[email protected]
Persistence of Disease Agents
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Menangle virus (porcine paramyxovirus)
Agent:
Family Paramyxoviridae, genus rubulovirus
Agent type:
Virus
Persistence and inactivation:
Very little experimental work has been conducted on this virus. It exists in only three
laboratories: EMAI, AAHL, and CTC (Atlanta). A reasonable assumption is that it has similar
properties to Newcastle disease virus, although limited field experience suggests MV may be
more fragile in the environment. Sentinel eight-week-old pigs introduced seven days after
depopulation failed to contract the disease.
The mode of transmission between pigs is unknown, but infection from faecal or urinary
contamination is considered to be more likely than by respiratory aerosols. There is no evidence
of a persistent shedding of the virus by infected animals. In the only cases of Menangle virus
reported to date, eradication was achieved using serological testing and segregation.
The virus is rapidly inactivated at low pH (Kirkland pers comm, Kirkland et al 2001).
Conclusions:
•
There is insufficient information available to make detailed recommendations on the
disposal or disinfection of animal carcases or products exposed to Menangle virus.
References:
Kirkland, P.D., Love, R.J., Philbey, A.W., Ross, A.D., Davis, R.J., & Hart, K.G. 2001,
‘Epidemiology and control of Menangle virus in pigs’, Aust Vet J, vol. 79, pp. 199-206.
Experts contacted:
Dr Peter Kirkland
NSW Agriculture
Elizabeth Macarthur Agricultural Institute
Woodbridge Rd
MENANGLE NSW 2568
P 02 4640 6331
[email protected]
Persistence of Disease Agents
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Newcastle disease
Agent:
Family Paramyxoviridae, genus paramyxovirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: Newcastle disease virus survives well in the environment. NDV may
persist for months at 8oC and for several days at 37oC (Beard and Hanson 1984).
NDV is readily inactivated by heating. Some studies have shown strain variation in heat lability,
unrelated to virulence (Arzey 1989). In nutrient broth, inactivation times at 56oC varied between
five minutes and six hours. All strains were inactivated at 100oC after 60 seconds (Beard and
Hanson 1984).
NDV is sensitive to a wide range of disinfectants, including chlorine-based chemicals and lipid
solvents (AFFA 2000).
Transmission of NDV between birds occurs through inhalation of droplets or ingestion of
material such as faeces, with the latter appearing to be the more important in the field (Alexander
1995). Rodents were shown to harbour the virus during one outbreak and were suggested as a
mechanism of spread in another (Arzey 1989).
The persistence and inactivation of NDV in poultry products has been thoroughly reviewed by
Arzey (1989).
Carcases and meat products: NDV is found at various times in most organs and tissues, the
distribution and concentration dependent on the virulence of the strain and on tissue tropism
(Hofstad 1951, Sinha et al 1952). Velogenic and mesogenic strains of NDV have been isolated
from carcases at slaughter (Lancaster and Alexander 1975). Gordon et al (1948) attributed one
third of the first 542 outbreaks of ND in England and Wales to feeding poultry waste to chickens.
Reid (1961) isolated NDV from up to two-thirds of frozen poultry imported into Britain that year.
Survival of the virus depends on its initial concentration, the ambient temperature and humidity,
and length of exposure (Foster and Thompson 1952, Olesiuk 1951). Reported survival times
include:
• 96 days on the skin and 134 days in bone marrow of eviscerated plucked carcases held at
2oC
• 160 days on the skin and 196 days in bone marrow of eviscerated unplucked carcases
held at 2oC
• greater than 300 days on skin and in bone marrow of both plucked and unplucked
eviscerated carcases held at -20oC (Asplin 1949)
• 270 days on skin and in meat at -14oC to -20oC for a mesogenic strain (Michalov et al
1967).
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Arzey (1989) discusses the heating requirements for inactivation of NDV in poultry meat
products, and the time/temperature combinations achieved in various commercial processes, at
some length. He notes that the thermostability data from studies using nutrient broth may not
apply in meat because of the protective environment offered by meats. One minute at 75oC is
suggested as sufficient for meat with low titre NDV. For waste foods fed to poultry, one minute
at 100oC may be required to remove all virus activity.
Wooley et al (1981) found that NDV survived for at least 96 hours (the entire test period) in
Lactobacillus-fermented food waste material held at 5oC, 10oC, 20oC, and 30oC. pH dropped as
low as 3.5 at 30oC.
Eggs and egg products: Newcastle disease virus has been isolated from eggs (e.g. Zagar and
Pomeroy 1950). Williams and Dillard (1968) found NDV in the membranes below the shell.
Dried egg has also been found to contain NDV (Alegren 1951). Eggs are therefore an infective
risk both from external faecal contamination and also from within the egg.
Arzey (1989) notes that NDV would be better protected against heat in the medium of a liquid
egg than it would in nutrient broth, in which the thermostability data of Foster and Thompson
(1957) were derived (see above). Gough (1973) isolated NDV from liquid egg that had been
heated to 64.4oC for 200 seconds. King (1991) reported that pasteurisation did not destroy NDV
in egg, serum or viral diagnostic agents.
Other products: NDV is found in the faeces of infected chickens. It has been reported to persist in
uncultivated manure for more than six months (Alexander et al 1985), and for 20 days in manure
heaped in mounds, where internal temperatures would have been higher (Zakomirdin 1963).
Conclusions:
•
Newcastle disease virus can survive in organic matter for months. However, the
virus is quite sensitive to heat and is inactivated by a wide range of disinfectants.
•
The ND virus is highly contagious. Biosecurity of potentially infective material is
very important.
•
Burning, rendering and burial are the preferred methods for disposal of infective
material.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2000, The importation of
non-viable eggs and products containing egg. Technical issues paper, December 2000, AFFA,
Canberra.
Alegren, A., 1951, ‘La peste Aviaire’, Bull Off Int Epizoot, vol. 35, p. 29, cited in AFFA 2000.
Alexander, D.J., Wilson, G.W.C., Russell, P.H., Lister, S.A., & Parsons, G. 1985, ‘Newcastle
disease outbreaks in fowl in Great Britain during 1984’, Vet Rec, vol. 117, pp. 429-434, cited in
Arzey 1987.
Alexander, D.J., 1995, ‘The epidemiology and control of avian influenza and Newcastle disease’,
J Comp Path, vol. 112, pp. 105-126.
Persistence of Disease Agents
Revised December 2003
31
Scott Williams Consulting Pty Ltd
Arzey, G. 1989, Technical Bulletin 42. Mechanisms of spread of Newcastle disease, Agdex
450/653, NSW Agriculture and Fisheries, Sydney.
Asplin, F.D. et al 1949, ‘Newcastle disease’, Proc of the XIV International Congress, London, pp.
361-371, cited in Arzey 1989.
Beard, C.W., & Hanson, R.P. 1984, ‘Newcastle disease’, In, Disease of Poultry, eighth edition,
eds Barnes, H.J., Calnek, B.W., Reid, W.M., & Yoder, H.W., Iowa State University Press, Ames,
Iowa, pp. 452-470, cited by AFFA 2000 and Arzey 1989.
Foster, N.M. & Thompson, C.H. 1957, ‘The comparative thermolability of four strains of
Newcastle disease virus of widely varying virulence’, Vet Med, vol. 52, pp. 119-121, cited in
AFFA 2000 and in Arzey 1989.
Gordon, R.F., Reid, J., & Asplin, F.D. 1948, ‘Newcastle disease in England and Wales. Official
report’, Eighth World’s Poultry Congress, Copenhagen, vol. 1, pp. 642-650, cited in Alexander
1995 and Guittet et al 1997.
Gough, R.E. 1973, ‘Thermostability of Newcastle disease virus in liquid whole egg’, Vet Rec, vol.
93, pp. 632-633, cited in AFFA 2000.
Guittet, M., Le Coq, H., & Picault, J.-P. 1997, ‘Risques de transmission de la maladie de
Newcastle par des produits avicoles contaminés’, Rev sci tech Off int Epiz, vol. 16, pp. 79-82.
Hofstad, M.S. 1951, ‘A quantitative study of Newcastle disease virus in tissues of infected
chickens’, Am J Vet Res, vol. 12, pp. 334-339, cited in Arzey 1989.
King, D.J. 1991, ‘Evaluation of different methods of inactivation of Newcastle disease virus and
avian influenza virus in egg fluids and serum’, Avian Diseases, vol. 35, pp. 505-514.
Lancaster, J.E., & Alexander, D.J. 1975, ‘Newcastle disease: virus and spread’, Monograph No.
11, Canada Department of Agriculture, Ottowa, cited in Arzey 1989.
Michalov, J. et al 1967, ‘Survival of mesogenic Newcastle disease virus I on wrapping materials
II in refrigerated broiler carcases’, Vet Bull, abstract p. 1424, cited in Arzey 1989.
Olesiuk, O.M. 1951, ‘Influence of environmental factors on viability of Newcastle disease virus’,
Am J Vet Res, vol. 12, pp. 152-155, cited in Arzey 1989.
Reid, J. 1961, ‘The control of Newcastle disease in Great Britain’, Br vet J, vol. 117, pp. 275-288,
cited in Alexander 1995 and Guittet et al 1997.
Sinha, S.K. et al 1952, ‘Comparison of the tropisms of six strains of Newcastle disease virus in
chicken following aerosol infection’, J Inf Dis, vol. 91, pp. 276-282, cited in Arzey 1989.
Williams, J.E., & Dilliard, L.H. 1968, ‘Penetration patterns of Mycoplasma gallisepticum and
Newcastle disease virus through the outer structures of chicken eggs’, Avian Dis, vol. 12, pp. 650657, cited in AFFA 2000.
Persistence of Disease Agents
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Scott Williams Consulting Pty Ltd
Wooley, R.E., Gilbert, J.P. Whitehead, W.K, Shotts, E.B., & Dobbins, C.N. 1981, ‘Survival of
viruses in fermented edible waste material’, Am J vet Res, vol. 42, pp. 87-90.
Zakomirdin, A.A. 1963, ‘Biothermal disinfection of manure from poultry with Newcastle disease
and infectious laryngotracheitis’, Vet Bull, abstract p. 1985, cited in Arzey 1989.
Zagar, S.L., & Pomeroy, B.S. 1950, ‘The effects of commercial living Newcastle disease virus
vaccines’, Am J Vet Res, vol. 11, pp. 272-277, cited in AFFA 2000.
Persistence of Disease Agents
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Scott Williams Consulting Pty Ltd
Nipah virus
Agent:
Family Paramyxoviridae, genus megamyxovirus (proposed name)
Agent type:
Virus
Persistence and inactivation:
Work at AAHL has shown that pigs can become infected with Nipah virus via either the oral
route or by parenteral inoculation, and that the virus is excreted via oronasal routes. Experience
in Malaysia has suggested urine, saliva, pharyngeal and bronchial secretions and also semen may
be involved in transmission. It is also suspected that dogs and cats may have become infected
and/or mechanically transferred the infection from pig trucks (Mohd Nor et al 2000).
Little information is available on the survival and inactivation properties of this virus (Daniels,
Kirkland pers comm). Malaysian authorities were approached to fund some work on the virus at
AAHL but declined, and it is doubtful much work has been done elsewhere.
Conclusions:
•
There is insufficient information available to make detailed recommendations on the
disposal or disinfection of animal carcases or products exposed to Nipah virus.
References:
Mohd Nor, M.N., Gan, C.H, & Ong, B.L. 2000, ‘Nipah virus infection in peninsular Malaysia’,
Rev sci tech Off int Epiz, vol. 19(1), pp. 160-165.
Experts contacted:
Dr Peter Daniels
CSIRO Livestock Industries
Australian Animal Health Laboratory
Private Bag 24
5 Portarlington Rd
Geelong Vic 3220
P 03 5227 5000
F 03 5227 5555
[email protected]
Dr Peter Kirkland
NSW Agriculture
Elizabeth Macarthur Agricultural Institute
Woodbridge Rd
MENANGLE NSW 2568
P 02 4640 6331
[email protected]
Persistence of Disease Agents
Revised December 2003
34
Scott Williams Consulting Pty Ltd
Peste des petits ruminants
Agent:
Family Paramyxoviridae, genus morbillivirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: Relatively little work has been done on the persistence of the peste des
petits ruminants virus outside the animal, although it is reported to have similar physico-chemical
characteristics to rinderpest virus (Rossiter and Taylor 1994). Taylor (1990 unpublished, cited in
Rossiter and Taylor 1994) reported on a strain of the virus from India2 with the following thermal
stability:
Temp (oC)
-20
4
37
56
Half-life
24.2 days
9.9 days
3.3 hours
2.2 minutes
Lefevre (1987) reported comparable data, estimating a half-life of around two hours at 37oC and
inactivation within one hour at 50oC. The virus studied by Taylor was stable between pH 5.85
and 9.5 and rapidly inactivated below pH 4.0 or above pH 11.0. Rossiter and Taylor (1994)
advise the disinfection of infected premises with lipid solvents or agents with low or high pH.
PPR is characterised by high viraemia and multiplication of the virus in a wide range of tissues.
PPRV is shed in ocular, nasal, and oral secretions and faeces of infected animals. Transmission is
mainly by inhalation of aerosols from close contact animals, and by nuzzling and licking.
Recently contaminated fomites may also be a source of infection (Rossiter and Taylor 1994).
Carcases and meat products: Viable virus can be recovered for at least eight days from lymph
nodes from carcases stored at 4oC (Lefevre 1987). No other references were found during this
review regarding the persistence of PPRV in carcases or meat products. It is reasonable to
assume that PPRV would be inactivated by the decline in pH accompanying rigor mortis. AQIS
(1999a) have concluded that the risk of introducing PPRV via sausage casings is probably low.
However, note should be made of several reports of persistence of rinderpest virus. Blackwell
(1984) cites recovery of active virus from carcases stored at 4oC for 30 days, and from carcases
aged for 24 hours and then kept at 4oC for 8 days. The same paper reports recovery of virus from
carcases buried for two months. Ezzatt et al (1970) have reported infectivity of meat refrigerated
for 7 days. Rinderpest virus may survive for several months in salted meat (Blaha 1989).
Although at least one outbreak of rinderpest has been attributed to the ingestion of fresh meat
(Rossiter 1994), the risk of this happening with sheep or goat meat is regarded as low
(MacDiarmid and Thompson 1997).
2
Geering et al note that there is some doubt whether the disease in India is caused by PPRV or rinderpest
virus, so these data should be treated with some caution.
Persistence of Disease Agents
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Scott Williams Consulting Pty Ltd
Milk and milk products: No primary references were found on the presence or persistence of
PPRV in milk. AUSVETPLAN (DPIE 1996a) is confusing in this regard, first extrapolating
information from rinderpest to suggest that PPRV may infect sheep or goat milk, then stating
unequivocally that the virus is present in milk.
If the virus is present in milk, prolonged survival is unlikely and pasteurisation should inactivate
the virus, but this is unconfirmed (AQIS 1999b). AUSVETPLAN (DPIE 1996a) stipulates the
heat treatment of milk for milk powder in the case of an outbreak.
Skins, hides and fibres: The presence of virus on the skin of infected animals, by either excretion
or external contamination, is highly likely. Rinderpest virus has been shown to rapidly lose
infectivity on adequately dried infected hides (Beaton 1932), and it is reasonable to extrapolate
this finding to PPRV. The virus is inactivated by the liming and pickling, so treated and partially
treated hides are not considered to pose an infective risk (AFFA 2001). However, salting of skins
is probably protective of the virus. In a draft assessment, AFFA (2001) has concluded that
unprocessed skins and hides from susceptible animals in countries with PPRV pose a high
quarantine risk.
AUSVETPLAN (DPIE 1996a) states that, in the case of an outbreak, skins, hides or fibres
deemed to be infected may need to be destroyed.
Semen/embryos: No primary references were found on the presence or persistence of PPRV in
semen or on embryos, although DPIE (1996a) states that the virus is present in semen and
embryos and is likely to be transmitted by them. The evidence on which this judgement is based
is unclear, but infection is clearly a risk given the isolation of virus from other secretions.
Faeces: PPRV may be found in the faeces of infected animals (Rossiter and Taylor 1994). No
specific references were found on its persistence. AUSVETPLAN (DPIE 1996a) states that in the
event of an outbreak, faeces should be buried.
Conclusions:
•
The peste des petits ruminants virus is a fragile virus which is unlikely to survive for
more than a few hours away from an animal host or animal products.
•
PPR is principally spread by the respiratory route, with close contact required
between infected and susceptible animals. However, recently contaminated fomites
may also be a source of infection.
•
Until more is known PPRV should be considered equivalent to Rinderpest virus that
can survive in meat for up to a month. Carcases from infected flocks/herds should
be rendered, burned or buried.
•
Milk should be pasteurised.
•
Salting of skins protects the virus from degradation. Fully processed skins and
scoured wool are a negligible disease risk.
•
Semen and embryos should not be used.
Persistence of Disease Agents
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References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999a, Importation of sausage casings into
Australia, import risk analysis, December 1999, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999b, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
Beaton, W.G. (1932), ‘Infectivity of shade dried hides from rinderpest infected cattle’, Indian J
Vet Sci Anim Husb, vol. 2, pp. 86-87, cited in Rossiter 1994.
Blackwell, J.H. 1984, ‘Foreign animal disease agent survival in animal products: Recent
developments’, J Am Vet Med Assoc, vol. 184, pp.674-679.
Blaha, T. 1989, in Applied veterinary epidemiology, Elsevier, Amsterdam, cited in AQIS 1999a.
DPIE (Department of Primary Industries and Energy) 1996a, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Peste des petits ruminants, DPIE, Canberra.
DPIE (Department of Primary Industries and Energy) 1996b, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Rinderpest, DPIE, Canberra.
Ezzatt, M.A.E., Kamel, J., Rofail, B., & Osman, F. 1970, ‘Keeping quality of rinderpest virus in
meat kept in the refrigerator’, Egyptian Vet Med Assoc J, vol. 30, pp. 997-1011, cited in DPIE
1996b.
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
Lefevre, P.C. 1987, ‘Peste des petits ruminants et infection bovipestique des ovins et caprins’, in
Etudes et synthèses de l’I.E.M.V.T., nombre 5, douzième édition, Institut d’Elevage et de
Médecine Vétérinaire des Pays Tropicaux, Maison Alfort, France, cited in Rossiter and Taylor
1994.
Rossiter, P.B. 1994, ‘Rinderpest’, in Infectious diseases of livestock, Vol II, eds. Coetzer, J.A.W.,
Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 735-757.
Rossiter, P.B., & Taylor, W.P 1994, ‘Peste des petits ruminants’, in Infectious diseases of
livestock, Vol II, eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press,
Cape Town, pp. 758-765.
Persistence of Disease Agents
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Scott Williams Consulting Pty Ltd
Rinderpest
Agent:
Family Paramyxoviridae, genus morbillivirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: Rinderpest virus is a relatively unstable virus. It retains infectivity for
only a few hours outside the host, except in very favourable conditions, when it may persist for up
to two to four days (Rossiter 1994). The virus shows greatest stability at pH 7.2-7.9, and is
rapidly inactivated outside pH range 5.6-9.6 (Geering et al 1995). It is killed quickly at
temperatures above 70oC (De Boer and Barber 1964) but small fractions of tissue-cultured virus
have survived heating at 56oC for 50 to 60 minutes and at 60oC for 30 minutes (Timoney et al
1988). Parsonson (1992) has cited further thermal stability data from Scott (1959):
Temp (oC)
Protein content of
medium
Time exposure
-70
-25
4
25
37
56
5% calf serum
5% calf serum
5% calf serum
0.125%
5% calf serum
5% calf serum
17 weeks
23 weeks
> 8 weeks
24 hours
48 hours
60 minutes
Initial titre
(log10TCID50/mL)
Titre reduction
No loss
No loss
Half life 9.2 days
2.14
0.5
Half life 165 minutes
Half life 3.5 minutes
Rinderpest virus is sensitive to light and UV radiation. It is inactivated by most disinfectants,
including phenol, chinosol, formalin, beta-propiolactone, trypsin, and 1M hydroxylamine. Strong
alkalis are the most effective agents (Timoney et al 1988).
Rinderpest virus is present in the ocular, nasal, oral, and vaginal secretions and faeces of infected
animals. It is transmitted by predominantly by direct contact and possibly by aerosols over short
distances or even by windborne spread (Rossiter 1994).
Carcases and meat products: Susceptibility to decreases in pH post mortem would suggest that
rinderpest virus does not generally survive long in carcases. Rossiter (1994), citing Curasson
(1932), states that the virus is inactivated by carcase decomposition within one to three days.
However, Blackwell (1984) reported recovery of active virus from carcases stored at 4oC for 30
days, and from carcases aged for 24 hours and then kept at 4oC for 8 days. The same paper
reports recovery of virus from carcases buried for two months. The virus has also been recovered
from meat frozen prior to rigor mortis (Blaha 1989). Ezzatt et al (1970) have reported infectivity
of meat refrigerated for 7 days. Outbreaks of rinderpest have been attributed to the ingestion of
fresh meat (Rossiter 1994).
Rinderpest virus may survive for several months in salted meat (Blaha 1989). No other reports
were found during this review or by AFFA (2001a) on the persistence or inactivation of
rinderpest virus in meat products.
Milk and milk products: No primary references were found on the presence or persistence of
rinderpest virus in milk. DPIE (1996) and AQIS (1999) cite Plowright (1964) to state that virus
Persistence of Disease Agents
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Scott Williams Consulting Pty Ltd
can be present in milk from 1-2 days before clinical signs develop and for up to 45 days after
recovery. The same information is quoted by Blackwell (1984), citing another review. Later
review articles do not mention the presence of rinderpest virus in milk (Rossiter 1994, Timoney et
al 1988).
If the virus is present in milk, prolonged survival is unlikely and pasteurisation should cause
inactivation, but this is unconfirmed (AQIS 1999). AUSVETPLAN (DPIE 1996) stipulates the
heat treatment of milk for milk powder from restricted and control areas in the case of an
outbreak.
Skins, hides and fibres: The presence of virus on the skin of infected animals, by either secretion
or external contamination, is highly likely. Infectivity is lost rapidly from adequately dried
infected hides (Beaton 1932). The virus is inactivated by the liming and pickling, so treated and
partially treated hides are not considered to pose an infective risk AFFA (2001b). However,
salting of skins is probably protective of the virus. In a draft assessment, AFFA (2001b) has
concluded that unprocessed skins and hides from susceptible animals in countries with rinderpest
pose a high quarantine risk.
AUSVETPLAN (DPIE 1996) stipulates disinfection of skins, hides or fibres before removal from
restricted and control areas in the case of an outbreak.
Semen/embryos: No primary references were found on the presence or persistence of rinderpest
virus in semen or on embryos, although DPIE (1996) refers to “very early work” demonstrating
semen transmission. Little virus is excreted via the reproductive tract (Philpott 1993). The virus
is found in vaginal discharges of infected cows (Rossiter 1994). There appears to be a real but
low risk of infectivity of semen or embryos.
Faeces: Rinderpest virus may be found in the faeces of infected animals (Rossiter 1994). No
specific references were found on its persistence.
Conclusions:
•
Rinderpest virus is a fragile virus which is unlikely to survive for more than a few
hours away from an animal host or animal products. The virus is readily
inactivated by exposure to heat, UV light and a wide range of disinfectants.
•
Rinderpest is principally spread by the respiratory route, with close contact
required between infected and susceptible cattle. However, recently contaminated
fomites may also be a source of infection. Oral infection has been reported.
•
Rinderpest virus can survive in meat for up to a month. Carcases from infected
herds should be rendered, burned or buried.
•
Milk from infected herds should be heat treated.
•
Salting of hides protects the virus from degradation. Fully processed skins are a
negligible transmission risk.
•
Semen and embryos should not be used.
Persistence of Disease Agents
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Scott Williams Consulting Pty Ltd
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001a, Generic import risk
analysis (IRA) for uncooked pig meat. Issues paper, January 2001, AFFA, Canberra.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001b, Import risk
analysis. Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
Beaton, W.G. (1932), ‘Infectivity of shade dried hides from rinderpest infected cattle’, Indian J
Vet Sci Anim Husb, vol. 2, pp. 86-87, cited in Rossiter 1994.
Blackwell, J.H. 1984, ‘Foreign animal disease agent survival in animal products: Recent
developments’, J Am Vet Med Assoc, vol. 184, pp.674-679.
Blaha, T. 1989, in Applied veterinary epidemiology, Elsevier, Amsterdam, 343 pp., cited in
MacDiarmid and Thompson 1997.
Curasson, G. 1932, La Peste Bovine, Vigot Freres, Paris, cited in Rossiter 1994.
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Rinderpest, DPIE, Canberra.
De Boer, C.J., & Barber, T.L. 1964, ‘pH and thermal stability of rinderpest virus’, Arch gesam
Virusfor, vol. 15, pp. 98-108, cited in DPIE 1996.
Ezzatt, M.A.E., Kamel, J., Rofail, B., & Osman, F. 1970, ‘Keeping quality of rinderpest virus in
meat kept in the refrigerator’, Egyptian Vet Med Assoc J, vol. 30, pp. 997-1011, cited in DPIE
1996.
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
MacDiarmid, S.C., & Thompson, E.J. 1997, ‘The potential risks to animal health from imported
sheep and goat meat’, Rev sci tech Off int Epiz, vol. 16(1), pp.45-56.
Parsonson, I.M 1992, Pathogen inactivation database documentation, consultancy for
Department of Primary Industries and Energy, Canberra.
Philpott, M. 1993, ‘The dangers of disease transmission by artificial insemination and embryo
transfer’, Br Vet J, vol. 149, pp. 339-345, cited in DPIE 1996.
Plowright, W. 1964, ‘Rinderpest virus’, Monographs in Virology, vol. 3, pp. 25-110, cited in
DPIE 1996 and AQIS 1999.
Rossiter, P.B. 1994, ‘Rinderpest’, in Infectious diseases of livestock, Vol II, eds. Coetzer, J.A.W.,
Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 735-757.
Persistence of Disease Agents
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Scott Williams Consulting Pty Ltd
Scott, G.R. 1952, ‘Heat-inactivation of rinderpest-infected bovine tissues’, Nature, vol. 184, pp.
1948-1949, cited in Parsonson 1992.
Timoney, J.F., Gillespie, J.H., Scott, F.W., & Barlough, J.E. (eds) 1988, ‘Rinderpest’, in Hagan
and Bruner’s microbiology and infectious diseases of domestic animals, eighth edition, Comstock
Publishing Associates, Ithaca.
Persistence of Disease Agents
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Scott Williams Consulting Pty Ltd
Picornaviridae
Foot-and-mouth disease
Agent:
Family Picornaviridae, genus aphthovirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: Foot-and-mouth disease has been thoroughly reviewed by Thomson
(1994). FMDV is very labile in acid and alkaline conditions. Donaldson (1987) states that
stability is greatest at 7.4-7.6, but with survival at 6.7-9.5 at below 4oC. Bachrach (1968)
reported that the time to reduce infectivity by 1 log10 at varying pH was:
pH
Time
5 1 second
6 1 minute
6.5 14 hours
10 14 hours
For differing temperatures, the time to reduce infectivity by 1 log10 was:
Temp
Time
4oC
20oC
37oC
49oC
61oC
1 week
11 days
21 hours
1 hour
3 seconds
(Woese 1960). Bengis (1997) notes that 69oC “appears to be the critical temperature”. FMDV is
stable almost indefinitely below 4oC (Donaldson 1987). FMDV is resistant to sunlight, but
sensitive to drying, with poor survival below a relative humidity of 55-60% (Thomson 1994).
AFFA (2001a) cites other reports of FMDV survival in the environment – for example, 50 days in
water; 26-200 days in soil, hay, sacking or straw; 345 days on one farm in California in 1924.
The relative lability of FMDV in alkaline conditions means that 2% NaOH or KOH and Na2CO3
can be used as cheap and effective disinfectants. Acids are also effective, but are more prone to
neutralisation by organic materials such as blood and faeces (Thomson 1994).
In viraemic animals, FMDV is found in all physiological fluids and therefore nearly all excretions
and secretions. The virus appears for up to four days prior to clinical signs. Infection takes place
via oral, respiratory and possibly venereal routes, while transmission is by direct contact, by
windborne spread, and by fomites. Different species showing varying susceptibility to these
modes (Thomson 1994).
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Carcases and meat products: The risk of FMDV infectivity in meat products has been reviewed
by Callis (1996), and in pig meat products specifically by AFFA (2001b) and Farez and Morley
(1997).
FMDV is distributed throughout the body. Survival of the virus post-mortem depends on the
stage of disease at time of slaughter, the strain of virus and environmental factors, especially
temperature and hydrogen ion concentration. The pH changes associated with rigor mortis are
sufficient to inactivate FMDV in muscle within 24 to 72 hours after slaughter. However, caution
in extrapolating findings between species is recommended by Bengis (1997), citing the case of
highly myotropic FMDV surviving beyond 72 hours in the carcase of impala, when the pH had
dropped to 5.6.
Refrigeration suspends the formation of acid, in which case the virus can survive for weeks or
months, especially in lymph nodes, blood clots, bone marrow and viscera (Henderson and
Brooksby 1948, Callis 1996). Cottral (1969) reported survival of the virus for 120 days in chilled
lymph nodes.
Gomes et al (1994) detected the O1 strain of FMDV in semimembranosus and longissimus dorsi
muscles and internal organs of sheep that were slaughtered during the febrile state. The muscles
did not reach a pH of less than 6. The virus was detectable both before and after maturation of
the carcase. It survived four months of freezing at -20oC. Virus was not detectable in sheep
slaughtered at 15 or 30 days post-infection.
The OIE International Animal Health Code recommends meat be heated to a core temperature of
at least 70oC for 30 minutes, but this may be insufficient to ensure complete inactivation.
Mincing generally helps to reduce FMDV survival and lactic-curing of salamis ensures
inactivation of any virus within a week. The virus can survive for long periods in high salt
concentrations, such as those found on sausage casings. Washing of infected intestines in 0.5% to
2.0% lactic or citric acid, or a citric acid buffer system (pH 5.3), for 8 to 10 hours is
recommended (MacDiarmid and Thompson 1997).
FMDV has also been shown to survive for up to:
• 60 days in beef held at 4oC in brine (Cottral et al 1960)
• 30 days in chilled pig lungs, stomach, tongue, intestine, 24 hours in chilled pig spleen,
liver and kidney, and 210 days in frozen pig organs (Savi et al 1962)
• 112 to 190 days in various hams and bacon (Dhenin et al 1980, McKercher et al 1987,
Mebus et al 1997)
• 42 days in Iberian pork loins (Mebus et al 1997)
• 56 days in pork sausages (Dhenin et al 1980)
• 250 days in processed pig intestinal casings (McKercher et al 1978)
• 7 days in pork salami (Panina et al 1989)
• 10 days in pork tongue and 1 day in muscle (Cottral 1969).
Milk and milk products: The high potential for transmission of FMDV via milk and milk products
has been reviewed by AQIS (1999a), Callis (1996) and Donaldson (1997). FMDV may be
excreted in milk before clinical signs are apparent and disappears with the development of
neutralising antibody (Callis 1996).
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The virus shows biphasic temperature and pH inactivation curves in which there is an initial rapid
phase, and then a more protracted phase of inactivation (Bachrach et al 1957). Some virus may
survive pasteurisation, probably due to its association with cell debris, although Donaldson
(1997) has argued that the risk posed by this small residue of virus, in terms of infecting calves or
pigs, is very small. Ultra high temperature (UHT) processing is sufficient to cause complete
inactivation (Cunliffe et al 1979). Heating of skim and full-cream milk to 80-90oC for 30 seconds
has been shown to reduce infectivity by 105.4 – 106.0 ID50 (Van Bekkum and de Leeuw 1978).
FMDV titre is reduced but not eliminated by processing of casein or caseinates, although after 30
days, infectivity could not be demonstrated (Callis 1996). Similarly, FMDV has been shown to
survive the processing of certain cheeses (e.g. cheddar made from milk that was not heat-treated,
for somewhere between 60 and 120 days at pH 5.0; camembert for one day after processing), but
infectivity disappears with ageing and ripening (Donaldson 1997).
In the opinion of Callis (1996), cheese production would be a good option of disposing of milk
collected during an outbreak. Milk from infected animals should not be fed back to livestock as
milk or other products (caseinate, whey, dry milk powder) unless the milk is UHT-processed.
Survival of FMDV in dried milk has been demonstrated with only a 2 log reduction at 100oC for
60 minutes (Nikitin and Vladimirov 1965). The effectiveness of heat treatment of viruses
generally increases with increasing water activity.
Skins, hides and fibres: In a draft assessment, AFFA (2001c) has identified hides and skins as
major potential hazards for the introduction of FMDV, both through surface contamination and
persistence in the skin itself.
FMDV virus has been recovered from dried hides for up to eight days, and from salted hides for
up to 352 days at 4oC (Gailiunis and Cottral 1967). The salt appears to be protective of the virus.
Sodium carbonate can be added to the salt to raise pH, but whilst the OIE International Animal
Health Code stipulates salting for 28 days with 2% added sodium carbonate to inactivate FMDV,
AFFA (2001c) points to the lack of information on the pH produced by this mixture. There is
also some confusion about whether the 2% is a proportion by weight of the hide or the salt.
The virus is inactivated by the high pH (>12.5) involved in liming and dehairing of skins and by
the low pH of pickling and tanning. Fully tanned or fully processed skins and hides (including
‘wet whites’ and ‘wet blues’) are therefore considered to pose a negligible risk. Disinfection with
acids or alkalis, low concentrations of formaldehyde, or 570g/m3 ethylene oxide gas at 52oC for
160 minutes will also be effective (AFFA 2001c).
FMDV has been isolated from greasy wool for up to 14 days after experimental contamination.
The virus survived for 7 weeks at 4oC, for 2 weeks at 18oC, and for 2 days at 37oC (McColl et al
1995).
Semen/embryos: The presence of FMDV in bull semen and its potential to infect cows during
artificial insemination has been demonstrated by Cottral et al (1968). FMDV has survived for a
month in frozen semen (Giefloff et al 1961). Virus may be shed for up to four days prior to
clinical signs, and for up to 42 days afterwards. Virus may be also be found in the semen of boars
before and after disease (Callis 1996, Sellers 1983). Recently, Bastros et al (1999) have
demonstrated the presence of FMDV in the semen and sheath of wild buffaloes.
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Callis (1996) summarised the work of several authors on the spread of FMDV by embryo
transfer, as follows:
• bovine, ovine and caprine embryos with intact zona pellucida were free of virus after
exposure to 106 plaque-forming units per mL of FMDV then washing according to
International Embryo Transfer Society standards;
• similar findings applied to embryos taken from acutely-infected stock and washed to
IETS standards;
• susceptible cows implanted with these embryos did not become infected;
• zona-pellucida intact bovine embryos resulting from insemination of a cow 90 days postinfection, with semen from a bull 40 days post-infection, and washed to IETS standards
were free of the virus;
• a similar study using sheep and goats resulted in virus-free embryos; but
• embryos without an intact zona pellucida did become infected.
These findings indicate a very low risk of transfer of FMDV by embryo transfer provided that
zona pellucida-intact embryos are used and are handled according to IETS recommendations.
AQIS (1999b) and Sutmoller and Wrathall (1997) have reached the same conclusion.
There is little information on swine embryos, although there is some indication that the virus may
be more prone to sticking to the zona pellucida than in bovine embryos (Callis 1996).
Faeces: Haas et al (1995), in a review of virus survival in liquid manure, quote several studies on
the survival of FMDV. Muller (1973) reported survival of FMDV for 21-103 days. Bøtner
(1990) reported survival times ranging from >14 weeks at 5oC to 1 hour at 55oC in pig slurry, and
between 5 weeks at 20oC and >60 minutes (inactivation not reached) at 55oC in cattle slurry.
Initial concentration of virus was 104.8 TCID50/50uL in each case. Eizenberger (unpublished,
cited in Haas et al 1995) demonstrated survival of FMDV in cattle slurry of 84 days at 4oC to 70
days at 17oC. AFFA (2001a) cites several figures for survival in manure, ranging from 6 to 42
days.
Conclusions:
•
FMD virus is a sensitive virus which, under normal conditions, does not survive for
more than a few days away from an animal host or animal products. However, the
virus can survive for months in a protected environment.
•
The FMD virus is sensitive to changes in pH, heat and drying.
•
Infection principally occurs by the respiratory and oral routes, with transmission by
direct contact, windborne spread and fomites. The virus is highly contagious.
Biosecurity of potentially infective material is very important.
•
Carcases should be buried or burned on-site. Burial, burning and rendering off-site
are less desirable options which should only be undertaken with a high level of
biological security during transportation.
•
Milk from infected herds should be heat treated. (Refer to the FMD AUSVETPLAN
strategy for more specific information).
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•
Salting of skins protects the virus from degradation. Unprocessed skins that may be
contaminated with FMD virus should be buried, burned or disinfected prior to
further processing. Fully processed skins are a negligible disease risk.
•
Wool that may be contaminated with FMD virus should be stored at 18ºC for four
weeks or sent for industrial scouring. Scoured wool poses a negligible disease risk.
•
Semen and embryos should be destroyed if collected less than one week prior to or
at any time after the first case of FMD in the herd.
References:
AFFA (Agriculture, Fisheries and Forestry – Australia) 2001a, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.2, Disease Strategy Foot-and-mouth disease, AFFA, Canberra.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001b, Generic import risk
analysis (IRA) for uncooked pig meat. Issues paper, January 2001, AFFA, Canberra.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001c, Import risk
analysis. Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999a, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999b, Import risk analysis report on the
importation of bovine semen and embryos from Argentina and Brazil into Australia, November
1999, AQIS, Canberra.
Bachrach, H.L. 1968, ‘Foot-and-mouth disease’, Ann Rev Microbiol, vol. 22, pp. 201-224, cited
in Thomson 1994.
Bachrach, H.L., Breese, S.S., Callis, J.J., Hess, W.R., & Patty, R.E. 1957, ‘Inactivation of footand-mouth disease virus by pH and temperature changes and by formaldehyde’, Proc Soc Exp
Biol Med, vol. 95, pp. 147-152, cited in Thomson 1994 and in Donaldson 1997.
Bastros, A.D.S., Bertschinger, H.J., Cordel, C., van Vuuren, C. deW.J., Keet, D., Bengis, R.G.,
Grobler, D.G., & Thomson, G.R. 1999, ‘Possibility of sexual transmission of foot-and-mouth
disease from African buffalo to cattle’, Vet Rec, vol. 145(3), pp.77-79.
Bengis, R.C. 1997, ‘Animal health risks associated with the transportation and utilisation of
wildlife products’, Rev sci tech Off int Epiz, vol. 16, pp. 104-110.
Bøtner, A. 1990, Modelstudier vederonde overlevelse af virus i gylle under traditionel opbevaring
og under udradning i biogasanlaeg. Research report, State Veterinaary Institute for Virus
Research, Lindholm, Denmark, cited in Haas et al 1995.
Callis, J.J. 1996, ‘Evaluation of the presence and risk of foot and mouth disease virus by
commodity in international trade’, Rev sci tech Off int Epiz, vol. 15(3), pp. 1075-1085.
Cottral, G.E., Cox, B.F., & Baldwin, D.E. 1960, ‘The survival of foot and mouth disease virus in
cured and uncured meat’, Am J vet Res, vol. 21(81), pp. 288-297, cited in Callis 1996.
Persistence of Disease Agents
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Scott Williams Consulting Pty Ltd
Cottral, G.E., Gailiunas, P., & Cox, B.F. 1968, ‘Foot-and-mouth disease virus in semen of bulls
and its transmission by artificial insemination’, Arch ges Virusforsch, vol. 23, pp. 362-377, cited
in Callis 1996.
Cottral, G.E. 1969, ‘Persistence of foot and mouth disease virus in animals, their products and the
environment’, Bull Off int Epiz, vol. 71(3-4), pp. 549-568, cited in Bengis 1997 and in Farez and
Morley 1997.
Cunliffe, H.R., Blackwell, J.H., Dors, R., & Walker, J.S. 1979, ‘Inactivation of milkborne footand-mouth disease virus at ultra-high temperatures’, J Food Protec, vol. 42, pp. 135-137, cited in
Donaldson 1997.
Dawe, P.S. 1974, ‘Viability of swine vesicular disease in carcases and faeces’, Vet Rec, vol. 94, p.
430.
Dhenin, L., Frouin, A., Gicquel, B., Bidard, J.P., & Labie, J. 1980, ‘Risks of disseminating FMD
virus by meat products : survival of FMD virus in dry sausage’, Bull Acad vet Fr, vol. 53, pp.
349-355, cited in Farez and Morley 1997.
Donaldson, A.I. 1987, ‘Foot-and-mouth disease: the principal features’, Irish vet J, vol. 41, pp.
325-327, cited in AFFA 2001a.
Donaldson, A.I. 1997, ‘Risks of spreading foot and mouth disease through milk and dairy
products’, Rev sci tech Off int Epiz, vol. 16(1), pp. 117-124.
Farez, S., & Morley, R.S. 1997, ‘Potential animal health hazards of pork and pork products’, Rev
sci tech Off int Epiz, vol. 16(1), pp. 65-78.
Gailiunas, P., & Cottral, G.E. 1967, ‘Survival of foot-and-mouth disease virus in bovine hides’,
Am J Vet Res, vol. 28, pp. 1047-53.
Giefloff, B.C.H., & Jacobsen, K.F. 1961, ‘On the survival of foot and mouth disease virus in
frozen bovine semen’, Acta Vet Scand, vol. 2, pp. 210-213, cited in Kahrs et al 1980.
Gloster, J., Hewson, H., Mackay, D., Garland, A., Donaldson, A., Mason, I., & Brown, R. 2001,
‘Spread of foot-and-mouth disease from the burning of animal carcases on open pyres’, Vet Rec,
vol. 148, pp. 585-586.
Gomes, I., Monteiro, E.M., Darsie, G.C., Ramalho, A.K., & Safon, M.C.F. 1994, ‘Survival of
foot-and-mouth disease virus O1 in carcasses of experimentally infected sheep, before an after the
maturation process’, Bol Centr Panam Fiebre Aftosa, vol. 60, pp. 49-59.
Haas, B., Ahl, R., Bohm, R., & Strauch, D. 1995, ‘Inactivation of viruses in liquid manure’, Rev
sci tech Off int Epiz, vol. 14(2), pp. 435-445.
Henderson, W.M. & Brooksby, J.B. 1948, ‘The survival of foot-and-mouth disease in meat and
offal’, J Hyg (Camb), vol. 46, pp. 394-402.
Persistence of Disease Agents
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Scott Williams Consulting Pty Ltd
Kahrs, R.F., Gibbs, E.P.J., & Larsen, R.E. 1980, ‘The search for viruses in bovine semen, a
review’, Theriogenol, vol. 14, pp. 151-165.
MacDiarmid, S.C., & Thompson, E.J. 1997, ‘The potential risks to animal health from imported
sheep and goat meat’, Rev sci tech Off int Epiz, vol. 16(1), pp.45-56.
McColl, K.A., Westbury, H.A., Kitching, R.P., & Lewis V.M., 1995, ‘The persistence of footand-mouth disease virus on wool, Aust vet J, vol. 72, pp. 286-292.
McKercher, P.D., Hess W.R., & Hamdy, F. 1978, ‘Residual viruses in pork products’, App
environ Microbiol, vol. 35(1), pp. 142-145, cited in Farez and Morley 1997.
McKercher, P.D., Yedloutschnig, R.J., Callis, J.J., Murphy, R., Panina, G.F., Civardi, A.,
Bugnetti, M., Fonn, E.H., Laddomada, A., Scarano, C., & Scatozza, F. 1987, ‘Survival of viruses
in ‘Prosciutto di Parma’ (Parma ham)’, Can Inst Food Sci Technol J, vol. 20(4), pp. 267-72, cited
in Farez and Morley 1997.
Mebus, C.A., Arias, M., Pineda, J.M., Tapiador, J., House, C., & Sanchez-Vizcaino, J.M. 1997,
‘Survival of several porcine viruses in different Spanish dry-cured meat products’, Food Chem,
vol. 59, pp. 555-559.
Muller, W. 1973, ‘Hygiene landwirtschaftlicher und kommunaler Abfallbeseitigungssystemen
[Hygiene of agricultural and municipal waste disposal systems]’, in Hohenheimer Arbeiten 69,
Ulmer, Stuttgart, Germany, cited in Haas et al 1995.
Nikitin Ye Ye & Vladimirov A.G ‘The survival of viruses in dried milk’ Veterinariya V42 1965
Panina, G.F., Civardi, A., Massirio, I., Scatozza, F., Baldini, P., & Palmia, F. 1989, ‘Survival of
foot and mouth disease virus in sausage meat products (Italian salami)’, Int J Food Microbiol,
vol. 8, pp. 141-148, cited in Farez and Morley 1997.
Parsonson, I.M. 1992, Pathogen inactivation database documentation, consultancy for
Department of Primary Industries and Energy, Canberra.
Savi, P., Baldelli, B., & Morozzi, A. 1962, ‘Présence et persistance du virus aphteux dans les
viandes de porcins et de bovins et dans leurs produits dérives’, Bull Off int Epiz, vol. 57, pp. 853890, cited in Farez and Morley 1997.
Sellers, R.F. 1983, ‘Transmission of viruses by artificial breeding techniques: a review’, J Royal
Soc Med, vol. 76, pp. 772-775.
Sutmoller, P., & Wrathall, A.E., ‘A quantitative assessment of the risk of transmission of footand-mouth disease, bluetongue and vesicular stomatitis by embryo transfer in cattle’, Prev vet
Med, vol. 32, pp. 111-132.
Thomson, G.R. 1994, ‘Foot-and-mouth disease’, in Infectious diseases of livestock, Vol II, eds.
Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 825852.
Persistence of Disease Agents
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Scott Williams Consulting Pty Ltd
Van Bekkum, J.G., & de Leeuw, P.W. 1978, ’Some aspects of foot-and-mouth disease virus in
milk’, Paper presented to the 6th International Congress of veterinary Science, La Plata,
Argentina, November 1978, cited in Donaldson 1997..
Woese, C. 1960, ‘Thermal inactivation of animal viruses’, Amer NY Acad Sci, vol. 83, pp. 741751, cited in Parsonson 1992.
Persistence of Disease Agents
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Experts contacted:
Dr Alex Donaldson
Institute for Animal Health
Pirbright Reference Laboratories
Ash Rd
Pirbright, Woking
Surrey GU24 0NF
UK
P +44 1483 232441
F +44 1483 232448
[email protected]
Persistence of Disease Agents
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Scott Williams Consulting Pty Ltd
Swine vesicular disease
Agent:
Family Picornaviridae, genus enterovirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: Swine vesicular disease virus is a very hardy agent. It is relatively stable
over a pH range of 2-12, and has survived for 164 days at pH 5.10 and pH 7.54 at 5oC. The virus
lost 6 logs of infectivity after 164 days at pH 2.88 and pH 10.14, and after 38 days at pH 1.92 and
pH 11.96 (Herniman et al 1973). SVDV is more resistant to heating and desiccation than FMD
virus (Geering et al 1995), and can withstand freezing, although it is inactivated at 69oC (Loxam
and Hedger 1983).
SVDV is resistant to many common disinfectants and detergents (DPIE 1996). Thomson (1994)
recommends the use of an alkali such as 1% sodium hydroxide for disinfection in the presence of
faeces or other organic matter.
The major source of virus in transmission is ruptured vesicles. SVDV is also excreted in the
faeces for 20 days or more, although this represents a less significant source of virus. Spread
occurs through direct contact, or indirectly via fomites or swill feeding. Airborne spread is not a
feature of the epidemiology. Infection requires a much smaller dose of virus through abraded
skin than it does to establish through oral, nasal or ocular routes (Geering et al 1995, Thomson
1994).
SVDV is preferentially found in the epithelium of the coronary band, tongue, snout and lips as
well as myocardium, tonsils and brain stem (Thomson 1994). McKercher et al (1974) have
demonstrated transmission of SVD to pigs fed infected meat.
Carcases and meat products: The survival of SVDV in carcases and meat products has been
comprehensively reviewed by Farez and Morley (1997). SVDV is resistant to the pH changes
accompanying rigor mortis (AFFA 2001a). It is stable in infected tissue kept at ambient or higher
temperatures for at least four months (Thomson 1994). Dawe (1974) reported that there was no
loss in infectivity of SVDV, 11 months after slaughter, in carcases frozen at -20oC. The skin
yielded 106 TCID (tissue culture infective doses) per gram, intercostal muscle 104 TCID, and rib
bone and kidney cortex 103 TCID, similar to titres detected at the beginning of the storage period.
Watson (1981) reported no drop in titre of SVDV in carcase material held for twelve months at
12-17oC.
A number of studies have looked at the persistence of SVDV in meat products, for example:
• up to 14 to 84 days in Iberian dry-cured loins and shoulders (within the commercial
curing period) (Mebus et al 1997)
• up to 470 days in Iberian and Serrano dry-cured hams (exceeding the commercial curing
period for Serrano ham) (Mebus et al 1997)
• at least six months in Parma hams (McKercher et al 1978)
• 200 days in dry salami, dry pepperoni sausage and intestinal casings (McKercher et al
1974)
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•
a minimum of 780 days in intestinal casings in another study (McKercher et al 1978).
Farez and Morley (1997) stated that “apparent thermal inactivation of SVD virus is obtained by
heating to at least 69oC”.
SVDV survives for many months in buried carcases (DPIE 1996).
Skins, hides and fibres: There is little primary information available on the persistence of SVDV
on skins. In a draft assessment, AFFA (2001b) has concluded that there is unlikely to be many
virus particles within the skin of an infected animal unless it is prepared early in an outbreak.
Given its hardy nature, however, there is a high likelihood that the virus would survive on skins
and hides during transport. Unprocessed or partially processed skins could not be relied on to be
free of virus. Only fully processed or fully tanned skins (including ‘wet whites’ and ‘wet blues’)
are considered to pose no quarantine threat.
Semen/embryos: SVDV has been found in the semen of infected boars from 2-3 days after
infection and 1-2 days before clinical signs. Attempts to establish infection in gilts by artificial
insemination with infected semen were generally unsuccessful. One pig receiving a large dose of
virus via the semen (108 pfu) did become infected, but so too did another pig in the room at the
time (McVicar et al 1977).
In the absence of other evidence, AQIS (2000) has concluded that transmission of SVDV via
artificial insemination is unlikely. Similarly, DPIE (1996) notes that transmission by infected
embryos is unlikely if proper procedures are followed.
Faeces: Dawe (1974) recovered SVDV from faeces for up to 138 days in ambient temperatures of
12-17oC. The temperature subsequently rose to 25oC and the virus was not recovered.
Conclusions:
•
The swine vescicular disease virus is a persistent virus which can survive away from
an animal host for months. It can survive a wide range of temperatures and pH,
and is resistant to commonly used disinfectants.
•
Infection principally occurs by the respiratory and oral routes, with spread by
direct contact, fomites or ingestion of infected material. The virus is highly
contagious. Biosecurity of potentially infective material is most important.
•
Carcases should be burned, rendered or buried. Burial must be in a manner that
precludes subsequent scavenging, at a site where there is no effluent discharge.
•
Unprocessed skins that may be contaminated with SVD virus should be buried,
burned or subjected to further processing. Fully processed skins are a negligible
disease risk.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001a, Generic import risk
analysis (IRA) for uncooked pig meat. Issues paper, January 2001, AFFA, Canberra.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001b, Import risk
analysis. Skins and hides. Draft report, August 2001, AFFA, Canberra.
Persistence of Disease Agents
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Scott Williams Consulting Pty Ltd
AQIS (Australian Quarantine and Inspection Service) 2000, Draft porcine semen risk analysis:
risk assessment and risk management options, March 2000, AQIS, Canberra.
Blackwell, J.H. 1987, ‘Viruses in products of food animals’, Dairy and food sanitation, vol. 7, pp.
398-401.
Dawe, P.S. 1974, ‘Viability of swine vesicular disease in carcases and faeces’, Vet Rec, vol. 94, p.
430.
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Swine Vesicular Disease, DPIE, Canberra.
Farez, S., & Morley, R.S. 1997, ‘Potential animal health hazards of pork and pork products’, Rev
sci tech Off int Epiz, vol. 16(1), pp. 65-78.
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
Herniman, K.A.J., Medhurst, P.M., Wilson, J.N., & Sellers, R.F. 1973, Vet Rec, vol. 93, pp. 620625, cited in Farez and Morley 1997 and Dawe 1974.
Loxam, J.G., & Hedger, R.S 1983, ‘Swine vesicular disease: clinical signs, epidemiology and
control’, Rev sci tech Off int Epiz, vol. 2, pp. 11-24, cited in DPIE 1996.
McKercher, P.D., Graves, J.H., Callis, J.J., & Carmichael, F. 1974, ‘Swine vesicular disease:
virus survival in pork products’, Proc Ann Meet US Anim Hlth Assoc, vol. 78, pp. 213a-216a.
McKercher, P.D., Hess, W.R., & Hamdy, F. 1978, ‘Residual viruses in pork products’, Appl
Environ Microbiol, vol. 35, pp. 142-145, cited in Blackwell 1987 and Farez and Morley 1997.
McVicar, J.W., Eisner, R.J., Johnson, L.A., & Pursel, V.G. 1977, ‘Foot and mouth disease and
swine vesicular disease viruses in boar semen’, Proc US An Hlth Assoc, vol. 81, pp. 221-230,
cited in AQIS 2000.
Mebus, C.A., Arias, M., Pineda, J.M., Tapiador, J., House, & Sanchez-Vizcaino, J.M. 1997,
‘Survival of several porcine viruses in different Spanish dry-cured meat products’, Food Chem,
vol. 59, pp. 555-559.
Thomson, G.R. 1994, ‘Swine vesicular disease’, in Infectious diseases of livestock, Vol II, eds.
Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 817819, cited in Farez and Morley 1997.
Watson, W.A. 1981, ‘Swine vesicular disease in Great Britain’, Can Vet J, vol. 22, pp. 195-200,
cited in AFFA 2001a.
Wooley, R.E., Gilbert, J.P., Whitehead, W.K., Shotts, E.B., & Dobbins, C.N. 1981, ‘Survival of
viruses in fermented edible waste material’, Am J vet Res, vol. 42, pp. 87-90.
Persistence of Disease Agents
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Teschen disease (porcine polioencephalomyelitis)
Agent:
Family Picornaviridae, genus enterovirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: Teschen disease virus is a relatively stable agent. Porcine enteroviruses
can survive more than 168 days in the environment at 15oC (Leman et al 1986). Derbyshire
(1989) stated that Teschen disease virus can remain infective for 15 minutes at 60oC and for
longer periods at 56oC, while Idriss (1983) reported inactivation of porcine enteroviruses by 8
weeks at 22oC and after 15-25 minutes at 56oC.
TDV survived for more than 168 days in water at 9-15oC, with chlorine at 0.1mg/L producing a 1
log reduction within two hours (Ottis 1976). TDV is stable between pH 2.8 and 9.5 but rapidly
inactivated outside this range (Derbyshire 1989). In a study involving ten common disinfectants,
only sodium hypochlorite and ethyl alcohol completely inactivated the virus (Derbyshire and
Arkell 1989).
TDV is spread via the oral-faecal route and probably also by fomites (Derbyshire 1989).
Carcases and meat products: No specific reports on the persistence or inactivation of TDV in
carcases or meat products were identified by this review, or in an issues paper by AFFA (2001a).
Skins, hides and fibres: AFFA (2001b), in a draft analysis, have concluded that processing or
partial processing of pig skins “should effectively eliminate the risk with porcine enteroviruses”.
Teschen disease virus is unlikely to survive the preliminary scalding of slaughtered pigs, or the
alkaline scouring, acid pickling or liming / dehairing of skins. Unprocessed skins would pose a
threat because of the stability of the agent in the environment.
Semen/embryos: Porcine enteroviruses have been found in semen (Phillips et al 1972), but while
TDV is likely to be present in semen during the viraemic stage, transmission via artificial
insemination is considered unlikely as viruses “do not productively infect early embryos”
(Thomson 1994).
Faeces: Derbyshire (1989) summarised studies showing that inactivation of the virus is more
rapid in aerated slurry – for example, 1 log decrease at 20oC in 2-4 days (aerated) vs 300 days at
5oC (unaerated). One strain of porcine enterovirus (T80 of serotype 2) was inactivated in pig
slurry by treatment with calcium hydroxide at pH 11.5 (Albrecht and Strauch 1980; Derbyshire
and Brown 1979; Lund and Nissen 1983).
Conclusions:
•
The Teschen disease virus is a persistent virus which can survive away from an
animal host for months. It can survive a wide range of temperatures and pH, and is
resistant to commonly used disinfectants.
•
Disease transmission occurs with an oral-faecal cycle.
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•
Meat and processed skins pose a negligible transmission risk.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001a, Generic import risk
analysis (IRA) for uncooked pig meat. Issues paper, January 2001, AFFA, Canberra.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001b, Import risk
analysis. Skins and hides. Draft report, August 2001, AFFA, Canberra.
Albrecht, H., & Strauch, D. 1980,’Das Umwalzbeluftungsverfahren (System Fuchs) zur
Behandlung von flussigen tierischen und kommunalen Abfallen. 10 Mitteilung: Die Wirkung der
Umwaltzbeluftung auf Viren der Picorna-, Reo- und Adenogruppe’, Berl Munch Tierarztl
Wochenschr, vol. 93, pp. 86-93, cited in Derbyshire 1989.
AQIS (Australian Quarantine and Inspection Service) 2000, Draft porcine semen risk analysis:
risk assessment and risk management options, March 2000, AQIS, Canberra.
Derbyshire, J.B. 1989, ‘Porcine enterovirus (polioencephalomyelitis)’, in Virus infections of
vertebrates, Vol 2 Virus infections of porcines, ed. M.B. Pensaert, Elsevier Science Publishers
B.V., Amsterdam, pp. 225-233.
Derbyshire, J.B., & Arkell, S. 1971, ‘The activity of some chemical disinfectants against Talfan
virus and porcine adenovirus’, Br Vet J, vol. 127, pp. 137-142, cited in Derbyshire 1989.
Derbyshire, J.B., & Brown, E.G. 1979, ‘The inactivation of viruses in cattle and pig slurry by
aeration or treatment with calcium hydroxide’, J Hyg, vol. 82, pp. 293-299, cited in Derbyshire
1989 and in AFFA 2001b.
Idriss, U.A. 1983, ‘Investigation of viruses isolated from pigs with gastroenteritis’, Veterinariya
(USSR), vol. 2, pp. 33-34, cited in AFFA 2001b.
Leman, A.D., Straw, B., Glock, R.D., Mengeling, W.L., Penny, R.H.C., & Scholl, E. 1986, in
Diseases of swine, 6th edition, Iowa State University Press, Iowa, cited in AFFA 2001b.
Lund, E., & Nissen, B. 1983, ‘The survival of enteroviruses in aerated and unaerated cattle and
pig slurry’, Agric Wastes, vol. 7, pp. 221-223, cited in Derbyshire 1989.
Ottis, K. 1976, Comparative studies on the persistence of viruses in drinking waters and surface
waters, 156 pp., 179 refs, cited in AFFA 2001b.
Phillips, R.M, Foley, C.W., & Lukert, P.D. 1972, ‘Isolation and characterisation of viruses from
semen and the reproductive tract of male swine’, J Am Vet Med Ass, vol. 161, pp. 1306-1316,
cited in AQIS 2000 and in Thomson 1994.
Thomson, G.R. 1994, ‘Teschen, Talfan and other reproductive disorders of pigs caused by
porcine enteroviruses’, in Infectious diseases of livestock, Vol II, eds. Coetzer, J.A.W., Thomson,
G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 813-816.
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Poxviridae
Lumpy skin disease
Agent:
Family Poxviridae, genus capripoxvirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: Lumpy skin disease virus is a stable agent, remaining viable in scab or
tissue for long periods. It is stable between pH 6.6 and 8.6. It has persisted for 5 days at 37oC in
this pH range without significant loss of titre (although AFFA (2001) note conflicting reports on
heat stability). LSDV has shown infectivity in dried skin lesions on the animal for at least 33
days, and 18 days in scrapings from dry lesions at room temperature (Weiss 1968). The closely
related sheep pox virus is inactivated by heating at 56oC and is susceptible to UV light (Geering
et al 1995).
LSDV is inactivated by a wide range of disinfectants, including the detergent SDS, ether, and
chloroform (Weiss 1968).
The transmission of LSD is not well elucidated (Davies 1991). Virus has been isolated from
nasal, ocular, and pharyngeal secretions, semen, milk and blood (Thomas and Mare 1945, Weiss
1968). Insect vectors appear to be the mains mode of transmission, with the stable fly Stomoxys
calcitrans likely to be implicated (Hunter and Wallace 2001).
Carcases and meat products: No evidence was found for the persistence of LSDV in the meat of
infected animals. AQIS (1999a) has determined that LSDV is unable to be transmitted via meat
or meat products.
Milk and milk products: LSDV may be found in the milk of infected animals (Davies 1991).
AQIS (1999b), using data from capripoxviruses in general, has determined that while high
temperature / short time pasteurisation is likely to substantially reduce the infectivity of
capripoxvirus in milk, it could not be relied on to guarantee total inactivation. There is some
evidence that conditions equivalent to the low temperature / long time method inactivate
capripoxvirus (62oC for 30 minutes), but the presence of fat, protein and other solids in milk may
protect the virus. The low pH of cheese may also be insufficient to inactivate the virus.
Skins, hides and fibres: LSDV has shown infectivity in dried skin lesions on the animal for at
least 33 days, and 18 days in scrapings from dry lesions at room temperature (Weiss 1968).
In a draft assessment, AFFA (2001) has concluded that unprocessed skins and hides from
susceptible animals in countries with LSD pose a high risk. Given the hardy nature of the virus, it
is highly likely to be found on skins from infected animals. Unprocessed or partially processed
skins could not be relied on to be free of virus. Only fully processed or fully tanned skins
(including ‘wet whites’ and ‘wet blues’) are considered to pose no quarantine threat.
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Semen/embryos: LSDV is found in semen for up to 22 days (Davies 1991). Kahrs et al (1980)
noted that the significance of semen in the transmission of the virus is not clear. No information
was found on the presence of LSDV in embryos.
Faeces: Reports on the presence or persistence of LSDV in faeces were not found. The virus
would be expected to be present as a contaminant.
Conclusions:
•
The lumpy skin disease virus is a persistent virus which can survive away from an
animal host for months. It can survive a wide range of temperatures and pH, but is
susceptible to commonly used disinfectants.
•
LSD is mainly spread by biting insects, with mechanical transmission and a cattleinsect-cattle cycle.
•
Unprocessed skins that may be contaminated with LSD virus should be buried,
burned or disinfected prior to further processing. Fully processed skins are a
negligible disease risk.
•
While meat poses a negligible transmission risk, blood and other by-products are
not. Therefore it is not practical to process carcasses at abattoirs.
•
Milk from infected herds should not be used. Burial and burning are the preferred
methods of disposal.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999a, Importation of sausage casings into
Australia, import risk analysis, December 1999, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999b, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
Davies, F.G. 1991, ‘Lumpy skin disease, an African capripox virus disease of cattle’, Br vet J,
vol. 147, pp. 489-503.
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
Hunter, P., & Wallace, D. 2001, ‘Lumpy skin disease in southern Africa: a review of the disease
and aspects of control’, J Sth Afr Vet Assoc, vol. 72, pp. 68-71.
Kahrs, R.F., Gibbs, E.P.J., & Larsen, R.E. 1980, ‘The search for viruses in bovine semen, a
review’, Theriogenology, vol. 14, pp. 151-165.
Thomas, A.D., & Mare, C.V.E. 1945, ‘Knopvelsiekte’, J Sth Afr Vet Med Assoc, vol. 16, pp. 3643, cited in Davies 1991.
Persistence of Disease Agents
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Weiss, J.A. 1968, ‘Lumpy skin disease’, in Virology monographs, vol. 3, Springer Verlag, New
York, pp. 112-282, cited in Davies 1991.
Experts contacted:
Dr Truuske Gerdes
Onderstepoort Veterinary Institute
Private bag X05
Onderstepoort 0110
SOUTH AFRICA
P +27 12 529 9114
F +27 12 529 9418
[email protected]
Persistence of Disease Agents
Revised December 2003
58
Scott Williams Consulting Pty Ltd
Sheep pox
Agent:
Family Poxviridae, genus capripoxvirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: Capripoxviruses are very stable agents in the environment. They are
sensitive to sunlight but may persist for up to six months in a cool and dark environment (Geering
1995, Kitching 2000). Sheep pox virus has been shown to persist in vesicular fluid for 2-3 years
stored at 0oC or -15oC (Drieux 1975).
Lumpy skin disease virus is reported to be stable over pH range 6.6-8.6 (Geering et al 1995).
Capripox viruses are acid labile and loss of infectivity occurs after exposure to pH 3 for 2 hours
(Dardiri 1978). A study using goat virus showed less lability to alkali conditions. There was only
a 1 log10 reduction at pH 8 (Datta and Soman 1991).
Ferreira (1973) reported the following reductions in infectivity at various time / temperature
combinations for SPV suspended in buffer at an initial concentration of 8 log10 TCID50/mL:
• 45oC, two hours, 2.3 log10
• 50oC, 30 minutes, 4 log10
• 50oC, one hour, 6 log10
• 55oC, 30 minutes, 4.6 log10
• 55oC, one hour, virus not detectable
• 60oC, 30 minutes, 5.6 log10
• 60oC, one hour, virus not detectable
• 65oC, five minutes, 5 log10
• 60oC, 30 minutes, virus not detectable.
Other authors have shown similar results, although there are some strain differences (AQIS
1999).
SPV is susceptible to a range of disinfectants and detergents, including lipid solvents and acids
(e.g. 2% hydrochloric acid, citric acid), 0.1-1.0% hypochlorite, aldehydes, alcohols and iodophors
(Dardiri 1978, DPIE 1996).
Sheep pox is transmitted by the respiratory route, infecting animals directly and indirectly via the
environment. All excretions and secretions from infected animals may contain virus. The virus is
also shed in scabs from lesions in the skin. Mechanical spread by insects is suspected but not
confirmed (Geering et al 1995, Munz and Dumbell 1994).
Carcases and meat products: Infection with sheep pox is followed by a viraemic phase, so SPV
could theoretically be present in muscle, although pH changes associated with rigor mortis would
be likely to inactivate the virus.
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Kirk (1981) states that sheep pox virus is known to occur in muscle tissue and lymph nodes but
the length of its survival there is unknown. He notes that if contamination were to occur during
slaughter, the virus could persist for a long time. MacDiarmid and Thompson (1997) observe that
such contamination has not been reported and conclude that meat is unlikely to pose a quarantine
risk for SPV.
Putrefaction destroys SPV (Blaha 1989). SPV does not infect species other than sheep and
sometimes goats, but mechanical transmission via insects may occur (Munz and Dumbell 1994).
These facts add weight to the argument that the meat and internal organs of infected sheep
carcases are unlikely to pose an infective risk. The skins and wool, however, do pose a problem.
AUSTVETPLAN (DPIE 1996) stipulates either burning or burial of infected carcasses. Spraying
with phenol, and strict control of vermin, may be required if a delay is likely between slaughter
and disposal.
Milk and milk products: The virus may be isolated in milk in the early stages of the disease when
the fever is evident (Davies 1991a), although infected milk plays a minor role in transmission of
the disease under natural conditions (Munz and Dumbell 1994).
Thermal inactivation data (see above) suggest that while high temperature / short time
pasteurisation is likely to reduce that capripoxvirus in milk significantly, it is possible that milk or
milk products could contain infective virus after treatment. Milk is more protective than the
buffer solutions in which the data were derived. The low pH of cheese could not be relied upon
for inactivation (AQIS 1999).
AUSTVETPLAN (DPIE 1996) stipulates that milk from suspect animals is to be destroyed by
heat, acid or buried.
Skins, hides and fibres: SPV can survive in wool or hair of recovered animals for up to three
months (Geering et al 1995). In a draft assessment, AFFA (2001) has concluded that unprocessed
skins and hides from susceptible animals in countries with SPV pose a high risk. Given the hardy
nature of the virus, it is highly likely to be found on the skin of infected animals. Unprocessed or
partially processed skins could not be relied on to be free of virus. Only fully processed or fully
tanned skins (including ‘wet whites’ and ‘wet blues’) are considered to pose no quarantine threat.
AUSTVETPLAN (DPIE 1996) stipulates that any skins, wool or fibre which may have been
contaminated must be burned or buried.
Semen/embryos: No references were found on the presence or persistence of SPV in semen or on
embryos. The closely-related lumpy skin disease virus has been found in semen up to 22 days
after infection (Davies 1991b). Orchitis has been reported in goats (Merza and Mushi 1990), and
intrauterine transmission can occur during cowpox (Mayr and Czerny 1990). AQIS (2000)
concludes that there is a risk of contamination of semen or embryos.
Faeces: No reports were found on the presence or persistence of LSDV in faeces. The virus
would be expected to be present as a contaminant. Manure is normally incinerated (Munz and
Dumbell 1994).
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Conclusions:
•
Sheep pox virus is a persistent virus which can survive away from an animal host
for months.
•
Infection mainly occurs by the respiratory route, either directly with close animal to
animal contact or indirectly from the environment.
•
Meat poses a negligible transmission risk, however the surface of fresh carcases may
be contaminated with sheep pox virus. For this reason, fresh carcases and byproducts should be buried or burned.
•
AUSVETPLAN requires that skins and wool which may have been contaminated
with sheep pox virus must be buried or burned. Fully processed skins are a
negligible disease risk.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 2000, An analysis of the disease risks, other
than scrapie, associated with the importation of ovine and caprine semen and embryos from
Canada, the United States of America and member states of the European Union, final report,
AQIS, Canberra.
Blaha, T. 1989, in Applied veterinary epidemiology, Elsevier, Amsterdam, 343 pp., cited in
MacDiarmid and Thompson 1997.
Dardiri, A.H. 1978, ‘Sheeppox’, in Reference manual on foreign animal diseases, USDA, Plum
Island, USA, pp. 140-146, cited in AFFA 2001.
Datta, S., & Soman, J.P. 1991, ‘Host range and physiochemical characterisation of ‘Ranchi’ strain
of goat-pox virus’, Indian J Anim Sci, vol. 61, pp. 955-957, cited in AQIS 1999.
Davies, F.G. 1991a, ‘Lumpy skin disease of cattle: a growing problem in Africa and the near
east’, World Animal Review, vol. 68, pp. 37-42, cited in DPIE 1996.
Davies, F.G. 1991b, ‘Lumpy skin disease, an African capripox virus disease of cattle’, Br vet J,
vol. 147, pp. 489-503.
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Sheep and Goat Pox, DPIE, Canberra.
Drieux, H. 1975, ‘Résistances des virus dans les produits d’origine animale’, Commission des
Communautés Européennes, Information internes sur I’agriculture, no. 165, cited by Kirk
(1981).
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Ferriera, C., 1973 ‘Comportamento do virus da variola ovina sob a accao de alguns agentes
fisicos e quimicos. [Effects of physical and chemical agents on sheep pox virus], Anais da escola
superior de medicina veterinaria, vol. 15, pp. 7-40, cited in AQIS 1999 and AFFA 2001.
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
Kirk, I.G.A. 1981, ‘Meat products as potential source for introduction of exotic animal diseases’,
Communication (Agriculture Canada, Food Production and Inspection Branch), no.28, pp. 4-7.
Kitching, R.P., 2000 ‘Sheep pox’, in Diseases of sheep, third edition, eds. Martin, W.B., &
Aitken, I.D., Blackwell Science, Oxford.
MacDiarmid, S.C., & Thompson, E.J. 1997, ‘The potential risks to animal health from imported
sheep and goat meat’, Rev sci tech Off int Epiz, vol. 16(1), pp.45-56.
Mayr, A., & Czerny, C.P. 1990, ‘Cowpox virus’, in Virus infections of vertebrates. Vol 3 Virus
infections of ruminants, eds. Dinter, Z., & Morein, B., Elsevier Science Publishers B.V.,
Amsterdam, pp. 9-16, cited in AQIS 2000.
Merza, M., & Mushi, E.Z. 1990, ‘Sheep pox virus’, in Virus infections of vertebrates. Vol 3 Virus
infections of ruminants, eds. Dinter, Z., & Morein, B., Elsevier Science Publishers B.V.,
Amsterdam, pp. 43-48, cited in AQIS 2000.
Munz, E., & Dumbell, K. 1994, ‘Sheeppox and goatpox’, in Infectious diseases of livestock, Vol I,
eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp.
613-615.
Experts contacted:
Dr Truuske Gerdes
Onderstepoort Veterinary Institute
Private bag X05
Onderstepoort 0110
SOUTH AFRICA
P +27 12 529 9114
F +27 12 529 9418
[email protected]
Persistence of Disease Agents
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Scott Williams Consulting Pty Ltd
Reoviridae
African horse sickness
Agent:
Family Reoviridae, genus orbivirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: The characteristics of African horse sickness virus have been well
summarised by Coetzer and Erasmus (1994). AHSV has been reported as persisting for:
• Ten minutes at 55-75oC
• Three months in a medium containing calf serum at 4oC (although infectivity is rapidly
lost at -25oC unless a stabiliser is added)
• At least six months at 4oC in saline containing 10% serum
• More than two years in putrid blood
• Twelve months in washed infected erythrocytes stored at 4oC.
The optimal pH for AHSV is 7.0-8.5, with the virus showing greater lability off the acid end of
this range. It is resistant to lipid solvents such as ether.
African horse sickness is spread by Culicoides midges. It is not contagious. Mechanical
transmission by biting flies may play a very minor role but the virus is sensitive to high
temperatures and desiccation (Coetzer and Erasmus 1994).
Carcases and meat products: No references were found on the persistence of AHSV in carcases or
meat products. Mellor (pers comm) believes that the work had not been done. The relative acid
lability of the virus suggests that it would be inactivated by the pH changes accompanying rigor
mortis.
Dogs have become infected by feeding on the carcases of infected animals (Van Rensburg 1981).
Skins, hides and fibres: The means of transmission of AHS suggests that the risk of skin, hides or
fibres being infective is almost zero. In a draft assessment, AFFA (2001) has concluded that
skins and hides pose a negligible quarantine risk for AHS.
Semen/embryos: No references were found on the presence of AHSV in semen.
Faeces: No references were found on the presence of AHSV in faeces.
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Conclusions:
•
The African horse sickness virus is a sensitive virus that is not able to survive for
any length of time away from an animal host.
•
African horse sickness is spread by Culicoides midges.
•
Special disposal precautions are not necessary, except for freshly dead animals,
which should be buried, burned or rendered.
•
Animal products pose a negligible transmission risk and no special disposal
measures are warranted.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
Coetzer, D.W., & Erasmus, B.J. 1994, ‘African horse sickness’, in Infectious diseases of
livestock, Vol I, eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press,
Cape Town, pp. 460-475.
Mellor, P.S. 1993, ‘African horse sickness: transmission and epidemiology’, Vet Res, vol. 24, pp.
199-212.
Van Rensburg, L.B.J., De Clerk, J., Groenewald, H.B., & Botha, W.S. 1981, ‘An outbreak of
African horse sickness in dogs’, J Sth Afr Vet Assoc, vol. 52, pp. 323-325, cited in Mellor 1993.
Experts contacted:
Dr Philip Mellor
Institute for Animal Health
Pirbright Reference Laboratories
Ash Rd
Pirbright, Woking
Surrey GU24 0NF
UK
P +44 1483 232441
F +44 1483 232448
[email protected]
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Bluetongue
Agent:
Family Reoviridae, genus orbivirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: Bluetongue virus is very stable in blood and tissue specimens at 20oC,
4oC, and -70oC but not at -20oC. In the absence of extraneous protein it is extremely unstable at
high temperatures. BTV is unstable below pH 6.5 and in the presence of disinfectants containing
acid, alkali, sodium hypochlorite and iodophors. It is relatively resistant to UV and gamma
radiation, and lipid solvents such as ether and chloroform (Verwoerd and Erasmus 1994).
Geering et al (1995) state that bluetongue is transmitted via Culicoides midges, and that there is
no transmission by direct contact between animals in the absence of the vector, nor by indirect
means. There is very little excretion or secretion of virus by infected animals. Transmission by
oral or aerosol means is highly unlikely to occur and products even from infected animals (with
the exception of semen) “can be disregarded as a source of infection” (Verwoerd and Erasmus
1994).
Carcases and meat products: No reports were uncovered regarding the persistence of bluetongue
in carcases or meat products. The acid lability of BTV would suggest inactivation of the virus
during post-mortem maturation of the carcase. AQIS (1999a) has determined that bluetongue “is
unable to be transmitted by meat or meat products” and AUSVETPLAN (DPIE 1996) states that
bluetongue does not survive outside living vectors or hosts.
Milk and milk products: No reports were uncovered of bluetongue virus being shed in milk.
AQIS (1999b) has determined that milk does not pose a quarantine threat for BTV.
Skins, hides and fibres: The means of transmission of BT suggests that the risk of skin, hides or
fibres being infective is almost zero. In a draft assessment, AFFA (2001) has concluded that
skins and hides pose a negligible quarantine risk for BT.
Semen/embryos: The risk of transmission of bluetongue via semen or embryos has been reviewed
by several authors (Roberts et al 1993, Sellers 1983).
BTV has been found in bull semen, during the period of viraemia, and is infective either by
natural or artificial insemination (Verwoerd and Erasmus 1994, Roberts et al 1993). The period
of viraemia varies to a generally accepted maximum of 50 days in cattle and 20 days in sheep
(Geering et al 1995). The number of organisms found in the semen of cattle is sufficient to infect
heifers, at least during a 25-day period from about day 10 of infection (Sellers 1983).
Roberts et al (1993) cite a series of papers by Luedke et al (Luedke et al 1977, 1983, Luedke and
Walton, 1981) describing persistent viraemia and shedding of BTV in semen by a bull infected in
utero. The bull was negative on serological testing and viraemic to 11 years of age.
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Bovine embryos with an intact zona pellucida are protected from BTV infection. Without a zona
pellucida, they are readily infected and will degenerate (Bowen et al 1982). The likelihood of
intact embryos carrying intracellular infection is therefore very low, although BTV could be
present in the embryonic environment in association with cellular blood elements. Washing
embryos ten times according the International Embryo Transfer Society manual (Stringfellow and
Seidel 1990) gives 95% confidence that the percentage of BTV-contaminated embryos from
viraemic donors would probably be not more than 1% (Sutmoller and Wrathall 1997).
The situation with sheep is less clear. Hare et al (1988) found BTV in the semen of rams but
were unable to transmit infection to ewes bred to these rams. These authors also reported nontransmission of infection by embryos washed ten times. Gilbert et al (1987) found that BTV
adhered to the zona pellucida of ovine embryos, and successfully transmitted infection via
embryos from viraemic ewes. However, the embryos were only washed three to four times. In
another study, ovine embryos exposed to BTV in vitro were not disinfected by ten washes, but the
viability of the embryos was not clear (Singh et al 1997).
Faeces: No reports of BTV appearing in faeces were uncovered during this review.
Conclusions:
•
Bluetongue is spread by Culicoides vectors. Transmission does not occur with direct
contact between infected and susceptible animals in the absence of the vector. Nor
does the virus spread by indirect means.
•
Carcases and animal products pose a negligible transmission risk and no special
disposal measures are warranted.
•
Embryos collected from infected donors should be washed according to IETS
standards before use. Semen collected from bulls during the infective phase should
not be used.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999a, Importation of sausage casings into
Australia, import risk analysis, December 1999, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999b, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 2000, An analysis of the disease risks, other
than scrapie, associated with the importation of ovine and caprine semen and embryos from
Canada, the United States of America and member states of the European Union, final report,
AQIS, Canberra.
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Bluetongue, DPIE, Canberra.
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Bowen R.A., Howard, T.H., & Pickett, B.W. 1982, ‘Interaction of bluetongue virus with
preimplantation embryos from mice and cattle’, Am J vet Res, vol. 43, pp. 1907-1911, cited in
Sudmoller and Wrathall 1997.
Gilbert, R.O., Coubrough, R.I., Weiss, K.E. 1987, ‘The transmission of bluetongue virus by
embryo transfer in sheep’, Theriogenology, vol. 27, pp. 527-540, cited in AQIS 2000.
Hare, W.C.D., Luedke, A.J., Thomas, F.C., Bowen, R.A., Singh, E.L., Eaglesome, M.D., Randall,
G.C.B., & Bielanski, A. 1988, ‘Nontransmission of bluetongue virus by embryos from
bluetongue virus-infected sheep’, Am J Vet Res, vol. 49, pp. 468-472.
Luedke, A.J, Jones, R.H., & Walton, T.E. 1977, ‘Overwintering mechanism for bluetongue virus:
biological recovery of latent virus from a bovine by bites of Culicoides varipennis’, Am J trop
Med Hyg, vol. 26, pp. 313-325, cited in Roberts et al 1993.
Luedke, A.J, Jochim, M.M., & Barber, T.L. 1983, ‘Serologic and virologic responses of a
Hereford bull persistently infected with bluetongue virus for eleven years’, Proc ann meet Am Ass
Vet Lab Diag, vol. 25, p. 115, cited in Roberts et al 1993.
Luedke, A.J, & Walton T.E. 1981, ‘Effect of natural breeding of heifers to a bluetongue virus
carrier bull’, Bovine Pract, vol. 16, pp. 96-100, cited in Roberts et al 1993.
Roberts, D.H., Lucas, M.H., & Bell, R.A. 1993, ‘Animal and animal product importation and the
assessment of risk from bluetongue and other ruminant orbiviruses’, Br vet J, vol. 149, pp. 87-99.
Sellers, R.F. 1983, ‘Transmission of viruses by artificial breeding techniques: a review’, J Royal
Soc Med, vol. 76, pp. 772-775.
Singh, E.L., Dulac, G.C, & Henderson, J.M. 1997, ‘Embryo transfer as a means of controlling the
transmission of viral infections. XV. Failure to transmit bluetongue virus through the transfer of
embryos from viraemic sheep donors’, Theriogenology, vol. 47, pp. 1205-1214, cited in AQIS
2000.
Stringfellow, D.A., & Seidel, S.M. 1990, Manual of the International Embryo Transfer Society,
second edition, IETS, Champaign, Illinois, cited in Sutmoller and Wrathall 1997.
Sutmoller, P., & Wrathall, A.E. 1997, ‘A quantitative assessment of the risk of transmission of
foot-and-mouth disease, bluetongue and vesicular stomatitis by embryo transfer in cattle’, Prev
vet Med, vol. 32, pp. 111-132.
Verwoerd, D.W., & Erasmus, B.J. 1994, ‘Bluetongue’, in Infectious diseases of livestock, Vol I,
eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp.
443-459.
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Equine encephalosis
Agent:
Family Reoviridae, genus orbivirus
Agent type:
Virus
Persistence and inactivation:
Little information could be found on the persistence and inactivation of this virus. Erasmus et al
(1970) were able to reduce infectivity in vitro with 0.5% trypsin, or with exposure to pH 3.0 for
one hour at 37oC. The virus was totally inactivated after 5 minutes at 60oC, with “considerable
loss” of infectivity at 56oC after one hour. It was resistant to chloroform.
Equine encephalosis virus is spread by arthropods, with Culicoides spp midges appearing to be an
important vector (Coetzer and Erasmus 1994).
Conclusions:
•
Equine encephalosis virus is spread by arthropod vectors.
•
Carcases and animal products derived from horses with equine encephalosis are
unlikely to pose a transmission risk for any significant period after death. No
special disposal measures are warranted.
References:
Coetzer, J.A.W., & Erasmus, B.J. 1994, ‘Equine encephalosis’, in Infectious diseases of livestock,
Vol II, eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape
Town, pp. 476-479.
Erasmus, B.J., Adelaar, T.F., Smit, J.D., Lecatsas, G., & Toms, T. 1970, ‘The isolation and
characterisation of equine encephalosis virus’, Bull Off int Epiz, vol. 74, pp. 781-789.
Persistence of Disease Agents
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Retroviridae
Jembrana disease
Agent:
Family Retroviridae, genus lentivirus
Agent type:
Virus
Persistence and inactivation:
Very little research of direct interest has been conducted. In summary, Jembrana disease virus
appears to be quite unstable, a characteristic consistent with other lentiviruses such as human
immunodeficiency virus (Wilcox, pers comm). The virus is sensitive to diethyl ether
(Kertayadnya et al 1996). In the absence of direct research, information on pulmonary
adenomatosis and maedi-visna viruses is likely to provide the best indicator of the properties of
Jembrana disease virus.
Jembrana disease is thought to be transmitted mechanically by biting insects and/or during mass
vaccination programs (Geering et al 1995).
Carcases and meat products: There is no published information on this aspect of the disease.
AQIS (1999a) has concluded that Jembrana disease “does not appear likely to be transmitted in
meat or meat products”. The virus survives for several months in infected spleens kept at -22oC
in Indonesia, although there appears to be substantial loss of infectivity (Wilcox pers comm). A
titration study reported by Kertayadnya et al (1996) showed a decline in virus particles from 108
to 102 ID50/mL in plasma stored at 4oC for 24 hours. Rapid freezing / thawing of plasma at -70oC
caused a reduction in titre from 108 to between 103 and 104 ID50/mL, a level which was
maintained during storage at -70oC for 2 months.
Milk and milk products: JDV has been found in the milk of infected cows during the viraemic
phase, and milk containing JDV has been shown to be capable of initiating disease (Soeharsono et
al 1995). There is limited information on how long the virus is excreted in milk. Research on
other lentiviruses suggests that normal pasteurisation procedures would inactivate the virus.
AQIS (1999b) has concluded that, in the absence of further information, dairy products must be
assumed to present a quarantine risk for JDV, and will only allow milk and milk products from
countries with Jembrana disease provided that the milk is pasteurised.
Skins, hides and fibres: JDV is unlikely to be stable on skins or fibres for a prolonged period. In
a draft assessment, AFFA (2001) has concluded that “the likelihood of transmission [of Jembrana
disease] via hides or skins would be negligible”.
Semen/embryos: JDV was not detected in the semen of experimentally infected bulls before or
immediately after the febrile phase by Soeharsono et al (1995). The risk of transmission by
semen appears to be low, but there is insufficient information to entirely rule it out.
Faeces: Excretion of virus in faeces is not mentioned in the literature as a feature of the disease.
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Conclusions:
•
The Jembrana disease virus is a fragile virus that is not able to survive for any
length of time away from an animal host.
•
Jembrana disease is transmitted mechanically by biting insects. It may also spread
iatrogenically, with mass vaccination or other intervention.
•
Animal carcases and skins pose a negligible transmission risk. No special disposal
measures are warranted.
•
Milk from infected herds should be pasteurised.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999a, Importation of sausage casings into
Australia, import risk analysis, December 1999, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999b, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
Kertayadnya, G., Soeharsono, S., Hartaningsih, N., & Wilcox, G.E. 1996, ‘The physicochemical
characteristics of a virus associated with Jembrana disease’, in Jembrana Disease and the Bovine
Lentiviruses, Proceedings of a Workshop 10-13 June Bali, Indonesia, eds. Wilcox, G.E.,
Soeharsono, S., Darma, D.M.N., & Copland, J.W., Australian Centre for International
Agricultural Research, Canberra, pp. 43-48.
Soeharsono, S., Wilcox, G.E., Dharma, D.M.N., Hartaningsih, N., Sulistyana, K., & Tenaya, M.
1995, ‘The transmission of Jembrana disease, a lentivirus disease of Bos javanicus cattle’,
Epidemiol Infection, vol.115, pp. 367-374.
Experts contacted:
Dr Graham Wilcox
Division of Veterinary and Biomedical Sciences
Murdoch University
South St
Murdoch 6150 WA
AUSTRALIA
P 08 9360 2448
[email protected]
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Maedi-visna
Agent:
Family Retroviridae, genus lentivirus
Agent type:
Virus
Persistence and inactivation:
Maedi-visna virus is readily inactivated by UV radiation, ethyl ether, chloroform, formaldehyde,
ethanol, phenol and trypsin (Petursson et al 1990, DeMartini pers comm). Virus infectivity is
relatively stable between pH 5.1 and 10. The following reductions in infectivity of virus
suspended in buffer at 19-21oC have been reported by Thormar (1965):
pH
Reduction in infectivity
Time
9.4
7.7
5.1
4.2
3.2
3.2
1 log10
1 log10
1 log10
1 log10
4 log10
5.5 log10
4 days
7 days
1 day
1.5 hours (maedi), 1 hour (visna)
0.5 hours (maedi)
0.5 hours (visna)
MVV is stable for months at -50oC and relatively resistant to repeated freezing and thawing
(Petursson et al 1990). One log10 (90%) reduction in infectivity was reported at 50oC in 1%
serum (pH 7.3-7.5) after 10 minutes, and a 5 log10 reduction at 56oC after 10 minutes (Thormar
1965).
MV is transmitted by direct contact, presumably via the respiratory route, but also through milk.
Indirect transmission through water contaminated with faeces may occur but is considered
unimportant (Geering et al 1995).
Carcases and meat products: This review did not uncover any specific studies on the persistence
of MVV in carcases or meat products. The lack of primary data was confirmed by DeMartini
(pers comm). AQIS (1999a) and MacDiarmid and Thompson (1997) have concluded that MVV
does not present a quarantine risk for transmission by meat or meat products.
Milk and milk products: The principal means of transmission of maedi-visna is via milk and
colostrum (Pepin et al 1998). AQIS (1999b) noted that the heat inactivation data of Thormar
(1965) should be interpreted with caution in regard to milk, because the virus is present in
monocyte/macrophage cells, although pasteurisation of goat milk at 56oC has been effective in
the inactivation of the closely related caprine arthritis-encephalitis virus. Pepin et al (1998)
recognised heating of ewe colostrum at 56oC for one hour, and pasteurisation of milk, as
components of a MV eradication program within a flock. AQIS (1999b) found no direct data on
the effect of HTST pasteurisation on milk containing MVV, nor could this review find any.
The data of Thormar (1965) suggested that the pH range of cheeses would inactivate the virus.
AQIS (1999b) has concluded that, in the absence of further information, dairy products must be
assumed to present a quarantine risk for JDV, and will only allow milk and milk products from
countries with maedi-visna provided that the milk is pasteurised.
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Skins, hides and fibres: There does not appear to be any primary data on the presence of MVV on
sheep skins or wool. MVV is present in all body fluids, so contamination could not be ruled out,
and there is horizontal transmission of MVV via respiratory secretions (Pepin et al 1998).
However the relative fragility of MVV suggests that the risk would be very low. In a draft
assessment, AFFA (2001) concluded that “the mode of transmission of [maedi-visna] is such that
hides and skin are unlikely to be a significant risk factor in such transmission”.
Semen/embryos: MVV has been detected in the semen of rams simultaneously infected with
MVV and ovine brucellosis (de la Concha-Bermejillo et al 1996), possibly as a result of
leucocytospermia. MVV was not found in uterine washes or washed embryos from infected ewes
(Woodall et al 1993). No studies have definitively demonstrated transmission of MVV by
venereal means (Pepin et al 1998). AQIS (2000) concluded that the risk of introducing MVV to
Australia via embryos was small, and via semen moderate.
Faeces: There has been a report of transmission of maedi-visna via drinking water contaminated
with faeces, but the oral/faecal route is not considered to be important part of the epidemiology
(Geering et al 1995).
Conclusions:
•
The maedi-visna virus is a fragile virus that is not able to survive for any length of
time away from an animal host.
•
Transmission requires close contact between infected and susceptible animals, with
the disease principally spread from dam to offspring via the colostrum or milk.
•
Animal carcases and skins pose a negligible transmission risk. No special disposal
measures are warranted.
•
Milk from infected herds should be pasteurised.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999a, Importation of sausage casings into
Australia, import risk analysis, December 1999, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999b, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 2000, An analysis of the disease risks, other
than scrapie, associated with the importation of ovine and caprine semen and embryos from
Canada, the United States of America and member states of the European Union, final report,
AQIS, Canberra.
de la Concha-Bermejillo, A., Magnus-Corral, S., Brodie, S.J., & DeMartini J.C. 1996, ‘Venereal
shedding of ovine lentivirus in infected rams’, Am J Vet Res, vol. 57, pp. 684-688, cited in Pepin
et al 1998.
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Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
MacDiarmid, S.C., & Thompson, E.J. 1997, ‘The potential risks to animal health from imported
sheep and goat meat’, Rev sci tech Off int Epiz, vol. 16(1), pp.45-56.
Pepin, M., Vitu, C., Russo, P., Mornex, J.-F., & Peterhans, E. 1998, ‘Maedi-visna virus infection
in sheep: a review’, Vet Res, vol. 29, pp. 341-367.
Petursson, G., Goergsson, G., & Palsson, P.A. 1990, ‘Maedi-visna virus’, in Virus infections of
vertebrates. Vol 3 Virus infections of ruminants, eds. Dinter, Z., & Morein, B., Elsevier Science
Publishers B.V., Amsterdam, pp. 431-440.
Thormar, H.1965, ‘A comparison of visna and maedi viruses; I – Physical, chemical and
biological properties’, Res Vet Sci, vol. 6, pp. 117-128, cited in Petursson et al 1990 and AQIS
1999.
Woodall, C.J., Mylne, M.J.A., McKelvey, W.A.C., & Watt, N.J. 1993, ‘Polymerase chain
reaction (PCR) as a novel method for investigating the transmission of maedi-visna virus (MVV)
by pre-implantation embryos’, in Proc 3rd international sheep veterinary congress, Edinburgh
June 1993, p. 232, cited in AQIS 2000.
Experts contacted:
Professor James DeMartini
College of Veterinary Medicine and Biomedical Sciences
Colorado State University
Fort Collins, Colorado 80523-1601
USA
P +1 970 491 7051
[email protected]
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Pulmonary adenomatosis
Agent:
Family Retroviridae
Agent type:
Virus
Persistence and inactivation:
Pulmonary adenomatosis virus has not been cultured in vitro, so there is very little primary
information on the survival and inactivation of the virus. It does not survive exposure and
desiccation for long (Verwoerd 1990).
The disease is thought to be transmitted by the respiratory route in aerosols (Geering et al 1995).
Carcases and meat products: No primary information is available on the survival of PAV in
carcases or meat products. Verwoerd (1990) states that the infection appears to remain localised
in the lung, and that “no evidence has been found to date of virus or viral antigens in any other
organ, including the blood.” Since then, Holland et al (1999) have identified proviral RNA and
DNA in peripheral blood mononuclear and other cells, so the absence of virus in the muscle could
not be dismissed. However, the virus is unlikely to survive long in the carcase given its relative
fragility. AQIS (1999a) has concluded that PAV does not present a quarantine risk for
transmission by meat or meat products.
Milk and milk products: Pulmonary adenomatosis virus is only known to be spread by respiratory
droplets (Verwoerd 1990). It is not considered to pose a quarantine hazard in dairy products
(AQIS 1999b).
Skins, hides and fibres: There does not appear to be any reported data on the presence of
pulmonary adenomatosis on sheep skins or wool. Presence of the virus on skin is unlikely, except
where contamination from infected aerosols has occurred. In a draft assessment, AFFA (2001)
concluded that “the mode of transmission of [pulmonary adenomatosis] is such that hides and
skin are unlikely to be a significant risk factor in such transmission”.
Semen/embryos: Parker et al (1998) demonstrated infection-free transfer of embryos from dams
in infected flocks, or from uninfected dams mated to an infected sire. This appears to be the only
study on venereal transmission of PAV. AQIS (2000), however, noted that “any concurrent
disease process in a [pulmonary adenomatosis] infected ram which increases the mononuclear
content of semen may lead to the production of [pulmonary adenomatosis] infected semen”. It
concluded that the risk of introducing PAV to Australia via embryos was small, and via semen
moderate.
Faeces: No reports were found to indicate the presence of PAV in faeces.
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Conclusions:
•
The pulmonary adenomatosis virus is a fragile virus that is not able to survive for
any length of time away from an animal host.
•
Disease spread occurs by the respiratory route, with close contact required between
infected and susceptible animals.
•
Animal carcases and products pose a negligible transmission risk. No special
disposal measures are warranted.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999a, Importation of sausage casings into
Australia, import risk analysis, December 1999, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999b, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 2000, An analysis of the disease risks, other
than scrapie, associated with the importation of ovine and caprine semen and embryos from
Canada, the United States of America and member states of the European Union, final report,
AQIS, Canberra.
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
Holland, M.J., Palmarini, M., Garcia-Goti, M., Gonzalez, L., McKendrick, I., de las Heras, M., &
Sharp, J.M. 1999, ‘Jaagsiekte retrovirus is widely distributed both in T and B lymphocytes and in
mononuclear phagocytes of sheep with naturally and experimentally acquired pulmonary
adenomatosis’, J Virol, vol. 73, pp. 4004-4008.
Parker, B.N.J., Wrathall, A.E., Saunders, R.W., Dawson, M., Done, S.H., Francis, P.G., Dexter,
I., & Bradley, R. 1998, ‘Prevention of transmission of sheep pulmonary adenomatosis by embryo
transfer’, Vet Rec, vol. 142, pp. 687-689.
Verwoerd, D.W. 1990, ‘Jaagsiekte (ovine pulmonary adenomatosis) virus’, in Virus infections of
vertebrates. Vol 3 Virus infections of ruminants, eds. Dinter, Z., & Morein, B., Elsevier Science
Publishers B.V., Amsterdam, pp. 453-463.
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Rhabdoviridae
Australian lyssavirus
Agent:
Family Rhabdovirus, genus lyssavirus
Agent type:
Virus
Persistence and inactivation:
No studies on the survival properties of Australian lyssavirus were uncovered during the course
of this review (the absence of any information was confirmed by McColl, pers comm and Field,
pers comm).
Conclusions:
•
Little is known about the persistence of Australian lyssavirus. It is likely to have
similar properties to rabies virus, but this has not been confirmed. Rabies is a
fragile virus that cannot survive for long away from an animal host and does not
persist in carcases or animal products.
References:
A review of bat lyssaviruses can be found in: McColl, K.A., Tordo, N., & Setien, A. 2000, ‘Bat
lyssavirus infections’, Rev sci tech Off int Epiz, vol. 19(1), pp.177-196.
Experts contacted:
Dr Ken McColl
CSIRO Livestock Industries
Australian Animal Health Laboratory
Private Bag 24
5 Portarlington Rd
GEELONG VIC 3220
P 03 5227 5000
F 03 5227 5555
[email protected]
Dr Hume Field
Department of Primary Industries
Locked Mailbag 4
MOOROOKA QLD 4105
P 07 3362 9566
[email protected]
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Rabies
Agent:
Family Rhabdovirus, genus lyssavirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: Rabies is a fragile virus, surviving only for short periods outside the host.
It is inactivated by heating to 56oC for 30 minutes, but can be preserved for years at temperatures
less than -60oC or by freeze drying and storing at -4oC (Swanepoel 1994). It is stable at pH 5-10
at 4oC, and labile at pH 3 in 30 minutes.
RV is susceptible to UV light and to lipid solvents (soapy water, ether, chloroform, acetone),
ethanol 45-75%, quaternary ammonium compounds (e.g. 0.2% cetrimide), and iodine
preparations 5-7% (AFFA 2001a, Bowen-Davies & Lowings 2000). However, the rate of
inactivation of RV by physical and chemical conditions is greatly modified by the stabilising
effects of polypeptides and other compounds (Wandeler pers comm, Michalski et al 1976).
Carcases and meat products: Transmission of rabies between animals through the ingestion of
contaminated tissues has been documented (see reviews of non-bite transmission of rabies by
Afshar 1979 and Swanepoel 1994). However, no descriptions of the survival of RV in carcases
or meat products were found during this review, and MacDiarmid and Thompson (1997) state that
rabies virus has never been isolated from meat.
Wandeler (pers comm) advised that the fragility of rabies viruses has “probably precluded
detailed studies of their inactivation in carcases”. There is an assumption that virus on the
exposed surfaces of the carcase would be inactivated within a few hours, whilst infectivity of
internal organs is lost within a few days in summer or many weeks in winter. Infectivity persists
longest in organs with a high ante-mortem load of virus (CNS, salivary glands, pancreas, and
adrenals). Persistence would be expected to be short-lived under most Australian conditions
given the heat lability of the virus (see AFFA 2001a).
Fooks (pers comm) has confirmed the paucity of data on persistence of rabies in carcases and
animal products, and speculated that the virus “would not survive for long (a few weeks)”. He
understood that rendering any carcase would completely inactivate the virus.
MacDiarmid and Thompson (1997) conclude that the probability of introducing rabies in meat or
meat products “must be considered remote”.
Milk and milk products: Transmission of rabies via suckling has been reported in several species
(Afshar 1979, Swanepoel 1994). These reports have been described as “rare and anecdotal” and
milk products are not considered to pose an import risk to Australia for rabies (AQIS 1999a).
Skins, hides and fibres: This review did not uncover any specific reports on the persistence of RV
on skins, hides or fibres. The risk of RV being present for any period of time is likely to be very
small, given the susceptibility of the virus to drying and UV light. In a draft report, AFFA
(2001b) has determined that “the mode of transmission of the disease is such that hides and skins
are unlikely to be a significant risk factor in the transmission [of rabies]”.
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Semen/embryos: There are no reports of rabies virus having been isolated from or transmitted by
semen in livestock. AQIS (1999b, 2000) have concluded that the risk of this occurring in cattle or
pigs is very low, although it notes that rabies virus has been isolated from the testis of a vampire
bat and that bovine serum can stabilise the infectivity of the virus during freezing and thawing.
Similarly, rabies virus has been demonstrated in one embryo, the uterus and ovaries of a skunk, so
contamination of bovine embryos cannot be entirely ruled out. AQIS requires only that semen
and embryos to be imported must come from donor animals showing no signs of clinical signs of
rabies during and for 15 days after collection.
Faeces: No reports were found of the presence of rabies virus in faeces.
Conclusions:
•
Rabies is a fragile virus that is readily inactivated by exposure to heat, UV light,
lipid solvents and commonly used disinfectants. The virus cannot survive for long
away from an animal host and does not persist in carcases or animal products.
•
Special disposal precautions are not necessary, except for freshly dead animals,
which should be buried, burned or rendered.
References:
Afshar, A. 1979, ‘A review of non-bite transmission of rabies virus infection’, Br Vet J, vol. 135,
pp. 142-148.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001a, Generic import risk
analysis (IRA) for uncooked pig meat. Issues paper, January 2001, AFFA, Canberra.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001b, Import risk
analysis. Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999a, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999b, Import risk analysis report on the
importation of bovine semen and embryos from Argentina and Brazil into Australia, November
1999, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 2000, Draft porcine semen risk analysis:
risk assessment and risk management options, March 2000, AQIS, Canberra.
Bowen-Davies, J., & Lowings, P. 2000, ‘Current perspectives on rabies. 2. Review of classical
rabies and its control’, In Practice, vol. 22(4), pp. 170-175.
MacDiarmid, S.C., & Thompson, E.J. 1997, ‘The potential risks to animal health from imported
sheep and goat meat’, Rev sci tech Off int Epiz, vol. 16(1), pp.45-56.
Michalski, F., Parks, N.F., Sokol, F., & Clark, H.F. 1976, ‘Thermal inactivation of rabies and
other rhabdoviruses: stabilization by the chelating agent ethylenediaminetetraacetic acid at
physiological temperatures’, Infect Immun, vol. 14(1), pp. 135-143.
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Swanepoel, R. 1994, ‘Rabies’, in Infectious diseases of livestock, Vol I, eds. Coetzer, J.A.W.,
Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 493-552.
Experts contacted:
Dr Alex Wandeler
Leader, Center of Expertise for Rabies
Canadian Food Inspection Agency
Ottawa Laboratory
ADRI-CPQP
3851 Fallowfield Rd
PO Box 11300
Ottawa, On. K2H 8P9
CANADA
P +1 613 228 6698
F +1 613 228 6661
[email protected]
Dr Tony Fooks
Veterinary Laboratories Agency
UK
[email protected]
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Vesicular stomatitis
Agent:
Family Rhabdovirus, genus vesiculovirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: As with other Rhabdoviridae, vesicular stomatitis virus is a relatively
fragile agent. Fong and Madin (1954) reported stability of VSV between pH 4-11.6, although
Wooley and Gilbert (unpublished, cited in Wooley et al 1981) inactivated VSV in 2 hours at pH
4-5. It is inactivated by temperatures over 50oC but can survive in soil at 4-6oC (Hanson and
McMillan 1990). VSV may be preserved for years at -60oC and by freeze-drying under vacuum
(Hanson 1981).
The virus survives for 3-4 days in infected saliva on buckets, feed racks and hay (Hanson 1952).
It is rapidly inactivated by all common disinfectants, including formalin, phenols and quaternary
ammonium compounds (Hanson and McMillan 1990).
The epidemiology of VS is poorly understood. Arthropods are probably important in
transmission, but viraemia is not a feature of the disease in pigs or horses. The virus is thought to
enter the animal only via insect bites or abrasions (Wilks 1994, Letchworth et al 1999). Lowgrade viraemia has been observed only in experimentally-infected cattle (Orrego et al 1987).
Carcases and meat products: VSV is not found in any edible tissues (Hanson 1981). No reports
were found on the presence of the virus in meat or meat products (see also AFFA 2001a).
Wooley et al (1981) reported the inactivation of VSV within two hours of being inoculated into a
can of food waste fermenting with Lactobacillus acidophilus. The virus could not be recovered
even at 5oC, suggesting the low pH (4-5) was contributing to inactivation.
Milk and milk products: No reports were found of VSV in milk. Hanson (1981) states that “the
virus, when present in milk, does not survive pasteurisation”. DPIE (1996) has interpreted this to
mean that VSV is present in raw milk, but AQIS (1999a) stated that an extensive review of the
literature had failed to reveal any original account of VSV being excreted or transmitted in milk
and concluded that milk did not present a quarantine hazard for the introduction of VSV.
However, vesicles of VS can be present on the teat of infected cows (Hanson 1981). These could
be expected to discharge virus into the milk.
Cliver (1973) found that VSV, artificially added at 105 plaque-forming units per 2 kg of prestarter samples, was not recoverable in curd before pressing during a stirred-curd cheddar
procedure.
Skins, hides and fibres: In a draft assessment, AFFA (2001) has concluded that, because of the
method of transmission and the unstable nature of the virus, spread of VSV via skins is unlikely
to occur. No specific quarantine measures are therefore considered necessary.
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Semen/embryos: This review did not uncover any reports of VSV having been isolated from or
transmitted by semen. AQIS (1999b, 2000) have concluded that the risk of transmission of VSV
by artificial insemination is very low, although equipment such as straws and containers could be
readily contaminated. Extrinsic contamination of uterine flushes by the virus may occur. VSV
adheres to the zona pellucida of bovine embryos and neither washing nor treatment with trypsin
can be relied upon for disinfection (Lauerman et al 1986, Stringfellow et al 1989).
Sutmoller and Wrathall (1997) noted that the brief viraemia observed in some cases of
experimentally-infected cattle may lend some suspicion to the contamination of embryos, but
concluded that the risk was close to zero.
Faeces: VSV does not appear in faeces or urine (Letchworth et al 1999).
Conclusions:
•
The vesicular stomatitis virus is a sensitive virus which, under normal conditions,
does not survive for more than a few days away from an animal host.
•
The epidemiology of VS is poorly understood. Animals are thought to become
infected only via insect bites or abrasions to the skin or mucus membranes.
•
Fresh carcases should be disposed of in a manner that prevents insect attack or
scavenging.
•
Meat and skins pose a negligible transmission risk. No special disposal measures
are warranted.
•
Milk from infected herds should be pasteurised.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001a, Generic import risk
analysis (IRA) for uncooked pig meat. Issues paper, January 2001, AFFA, Canberra.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001b, Import risk
analysis. Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999a, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999b, Import risk analysis report on the
importation of bovine semen and embryos from Argentina and Brazil into Australia, November
1999, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 2000, Draft porcine semen risk analysis:
risk assessment and risk management options, March 2000, AQIS, Canberra.
Cliver, D.O. 1973, ‘Cheddar cheese as a vehicle for viruses’, J Dairy Sci, vol. 56, pp. 1329-1331.
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Vesicular Stomatitis, DPIE, Canberra.
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Fong, J., & Madin, S.H. 1954, ‘Stability of VSV at varying pH concentrations’, Proc Soc Exp
Biol Med, vol. 86, pp. 676-678, cited in AFFA 2001a.
Hanson, R.P. 1952, ‘The natural history of vesicular stomatitis’, Bacteriological Reviews, vol. 16,
pp. 179-204, cited in Letchworth et al 1999.
Hanson, R.P. 1981, ‘Vesicular stomatitis’, in Virus diseases of food animals. Vol II Disease
monographs, ed. Gibbs, E.P.J., Academic Press, London, pp. 517-539.
Hanson, R.P, & McMillan, B. 1990, ‘Vesicular stomatitis virus’, in Virus infections of ruminants,
eds. Dinter, Z., & Morein, B., Elsevier Science Publishers B.V., Amsterdam, pp. 381-391.
Lauerman, L.H., Stringfellow, D.A., Sparling, P.H., & Kaub, L.M. 1986, ‘In vitro exposure of
preimplantation bovine embryos to vesicular stomatitis virus’, J Clin Microbiol, vol. 24, pp. 380383, cited in AQIS 1999b.
Letchworth, G.J., Rodriguez, L.L., & Barrera, J. del C. 1999, ‘Review. Vesicular stomatitis’, Vet
J, vol. 157, pp. 239-260.
Orrego, A., Arbelaez, G., & Roch, J. 1987, ‘Avances en las investigaciones sobre le estomatitis
vesicular en Colombia’, Min de Agric, Inst Colombiano Agropecuario, 1st edición, Bogota, cited
in Sutmoller and Wrathall 1997.
Stringfellow, D.A., Lauerman, L.H., & Thomson, M.S. 1989, ‘Trypsin treatment of bovine ova
after in vitro exposure to vesicular stomatitis virus’, Am J Vet Res, vol. 50, pp. 990-992, cited in
AQIS 1999b.
Sutmoller, P., & Wrathall, A.E. 1997, ‘A quantitative assessment of the risk of transmission of
foot-and-mouth disease, bluetongue and vesicular stomatitis by embryo transfer in cattle’, Prev
vet Med, vol. 32, pp. 111-132.
Wilks, C.R. 1994, ‘Vesicular stomatitis and other vesiculovirus infections’, in Infectious diseases
of livestock, Vol II, eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University
Press, Cape Town, pp. 563-566.
Wooley, R.E., Gilbert, J.P., Whitehead, W.K., Shotts, E.B., & Dobbins, C.N. 1981, ‘Survival of
viruses in fermented edible waste material’, Am J vet Res, vol. 42, pp. 87-90.
Experts contacted:
Dr Sabrina Swenson
Head, Bovine and Porcine Viruses Section
Diagnostic Virology Laboratory
National Veterinary Services Laboratories
1800 Dayton Rd
Ames, Iowa, 50010
USA
P +1 515 663 7551
F +1 515 663 7348
[email protected]
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Togaviridae
Getah virus disease
Agent:
Family Togaviridae, genus alphavirus, Semliki Forest antigenic complex
Agent type:
Virus
Persistence and inactivation:
Getah virus is spread by mosquitoes (Geering et al 1995). Disease can be induced experimentally
in horses by subcutaneous, intramuscular and intranasal inoculation, and has been induced in pigs
by intramuscular inoculation, but while spread by direct contact is conceivable it appears to be
unimportant in the field (Kono 1988). In mice, there is horizontal spread between littermates and
vertical offspring via milk (Kono 1988).
Infected horses have a short viraemia, with virus localising in a wide range of tissues including
lymph nodes, lung, spleen, liver and bone marrow (Kamada et al 1981). Sentsui and Kono
(1980) did not find infective Getah virus in faeces or urine of infected horses.
No references were found on the persistence of Getah virus in meat or other animal products.
The closely related Western, Eastern and Venezuelan equine encephalomyelitis viruses are
reported to be extremely fragile, disappearing from infected tissues within a few hours after death
(Radostits et al 2000).
Conclusions:
•
The Getah virus is a fragile virus that cannot survive for long away from an animal
host and does not persist in carcases or animal products.
•
Getah virus is spread by mosquitoes.
•
Carcases and animal products pose a negligible transmission risk. No special
disposal precautions are warranted.
References:
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
Kamada, M., Ando, Y., Fukunaga, Y., Imagawa, H., Wada, R., Kumanomido, T., Tabuchi, E.,
Hirasawa, K., & Akiyama, Y. 1981, ‘Studies on Getah virus – pathogenicity of the virus for
horses’, Bull Equine Res Inst (Jpn), no. 18, pp. 84-93.
Kono, Y. 1988, ‘Getah virus disease’, in The arboviruses: epidemiology and ecology. Vol III, ed.
Monath, T.P., CRC Press, Boca Raton, Florida, pp. 21-36.
Radostits, O.M., Gay, C.G., Blood, D.C., & Hinchcliff, K.W. (eds) 2000, Veterinary medicine,
ninth edition, W. B. Saunders Company Ltd, London.
Sentsui, H., & Kono, Y. 1980, ‘An epidemic of Getah virus infection among racehorses: isolation
of the virus’, Res vet Sci, vol. 29, pp. 157-161, cited in Kono 1988.
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Porcine reproductive and respiratory syndrome
Agent:
Family Togaviridae, genus arterivirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: The survival properties of porcine reproductive and respiratory syndrome
virus have been summarised by AFFA (2001a).
PRRSV appears to be most stable in the pH range 5.5-6.5 (Bloemraad et al 1994), with Benfield
et al (1992) reporting 90% inactivation of the virus outside the pH range 5-7.
Heat stability studies on the European strain of the virus have shown survival for at least 72 hours
at 4oC or -20oC, but 93% loss of infectivity after 72 hours at 25oC (Alstine et al 1993). The US
strain is stable for at least 18 months at -70oC, for at least one month at 4oC, and loses 50% of
viability after 12 hours. Complete inactivation was achieved after 48 hours at 37oC and after 45
minutes at 56oC (Benfield et al 1992).
Bloemraad et al (1994) found in culture medium at pH 7.5 the virus half life was:
• 140 hours at 4oC
• 20 hours at 21oC
• 3 hours at 37oC
• 6 minutes at 56oC
The half life was decreased by rapid changes in pH.
The virus persists in the environment for up to three weeks (Geering et al 1995). Transmission of
PRRS occurs via aerosols. Windborne spread is a feature of the disease. Virus has been isolated
from faeces, urine, and semen (Done et al 1996). Rodents do not appear to be a reservoir of the
disease (Hooper et al 1994).
Magar (pers comm) advised no direct work had been conducted on disposal methods for infected
carcases or animal products, and none were uncovered during this review.
Carcases and meat products: After infection, PRRS probably spreads during viraemia to various
organs and tissues (Magar et al 1995).
Magar et al (1995) looked for PRRSV in the tissues of pigs who were either experimentally
infected or who came from infected herds. In the first group, virus was detected at 7 days postinoculation lungs, tonsils, lymph nodes and muscle tissue. Virus was not detected in muscle at 14
days but was found in lungs and tonsils.
No virus could be found in the second group, seropositive pigs from known infected herds, in
pools of tissue from 44 pigs. (Virus was found only in one lymph node pool, and it could not be
grown in alveolar macrophages.) The time since the sampled pigs had been exposed to the virus
was not known, but was probably weeks to months. The authors concluded that it was unlikely
that the disease would be transmitted by pork.
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European and American strains of PRRSV have been demonstrated in pooled samples of ham
muscle and bone marrow in pigs slaughtered six days post-infection (Frey et al 1995). The
pooled muscle bone marrow samples retained infectivity for several weeks when at 4 0C and at
least one month when stored at -20 0C.
A study by Larochelle and Magar (1997), using a sensitive PCR assay, failed to detect PRRSV in
438 muscle tissue homogenates from 73 different lots of packaged meat collected over seven
months.
In a study commissioned by AQIS at Lelystad ID-DLO (AFFA 2001a), European and American
strains of PRRSV were successfully transmitted to receiver pigs by feeding muscle tissue from
experimentally-infected pigs. Infection was confirmed by virus isolation and antibody detection
three weeks after feeding.
Skins, hides and fibres: No specific references were found on this aspect of the disease. In a draft
assessment, AFFA (2001b) determined that these products do not pose a quarantine risk for PRRS.
Semen/embryos: PRRSV has been detected in semen from three days, and for up to 92 days after
infection in some pigs (Christopher-Hennings 1995).
Experimental transmission of PRRSV to gilts in semen was recently achieved by Gradil et al
(1996). Seroconversion was demonstrated in gilts inseminated with semen from boars that had
been inoculated with PRRSV.
Faeces: PRRSV has been isolated from faeces of infected pigs (Done et al 1996).
Conclusions:
•
The PRRS virus is a sensitive virus that can survive in the environment for up to
three weeks, and longer in chilled and frozen meat products.
•
Transmission of PRRS occurs principally by respiratory aerosol, with windborne
spread a feature of the disease. Oral infection can also occur.
•
Fresh carcases should be disposed of in a manner that prevents scavenging. Meat is
a risk and should be cooked for at least 700C for 11 minutes. Fresh carcases should
be processed to at least this time/temperature or be buried, rendered or burnt.
•
Skins pose a negligible transmission risk, and no special disposal precautions are
warranted.
•
Semen collected from infected boars should not be used.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001a, Generic import risk
analysis (IRA) for uncooked pig meat. Issues paper, January 2001, AFFA, Canberra.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001b, Import risk
analysis. Skins and hides. Draft report, August 2001, AFFA, Canberra.
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Alstine, W.G. van, Kanitz, C.L., & Stevenson G.W. 1993. ‘Time and temperature survivability of
PRRS is serum and tissues’, J Vet Diagn Invest, vol. 5, pp. 621-622, cited in AFFA 2001a.
AQIS (Australian Quarantine and Inspection Service) 2000, Draft porcine semen risk analysis:
risk assessment and risk management options, March 2000, AQIS, Canberra.
Benfield, D.A., Harris, L., Nelson, E.A., Collins, J.E., Chladek, D.W., Christianson, W.T.,
Morrison, R., & Gorcyca, D. 1992, ‘Properties of a swine infertility and respiratory syndrome
virus (AT VR-2332) isolated in the United States’, in Proceedings of the International
Symposium on SIRS/ PRRS/ PEARS, Minnesota, USA, p.6, cited in AFFA 2001a.
Bloemraad, M., de Kluijver, E.P., Petersen, A., Burkhardt, G E., & Wensvoort, G. 1994, ‘Porcine
reproductive and respiratory syndrome: temperature and pH stability of Lelystad virus and its
survival in tissue specimens from viraemic pigs’, Vet Microbiol, vol. 42, pp. 361-371, cited in
AFFA 2001a.
Christopher-Hennings, J., Nelson, E.A., Nelson, J.K., Hines, R. J., Swenson, S.L., Hill, H.T.,
Katz, J.B., Yaeger, M.J., Chase, C.C.L., & Benfield, D. 1995, ‘Persistence of porcine
reproductive and respiratory syndrome virus in serum and semen of adult boars’, J Vet Diagn
Invest, vol. 7, pp. 456-464, cited in AQIS 2000.
Done, S.H., Paton, D.J., White, M.E.C. 1996, ‘Porcine reproductive and respiratory syndrome
(PRRS): A review, with emphasis on pathological, virological and diagnostic aspects’, Br Vet J,
vol. 152, pp. 153 cited in AQIS 1999.
Frey, M.L., Mengeling, W.L., Lager, K.M., Landgraf, J.G. & Eernisse, K.A. 1995, ’Survival of
the virus of porcine reproductive and respiratory syndrome (PRRS) in swine carcasses’, in
National (US) Veterinary Services Laboratories Development Project Final Report, pp 1-9, cited
in AFFA 2001a.
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
Gradil, C., Dubuc, C., Eaglesome, M.D. 1996, ‘Porcine reproductive and respiratory syndrome
virus: seminal transmission’, Vet Rec, vol. 138, pp. 521-522.
Hooper, C.C., Alstine, W.G., Stevenson, G.W., Kanitz, C.L. & van Alstine, W.G. 1994, ‘Mice
and rats (laboratory and feral) are not a reservoir for PRRS virus’, J Vet Diagnostic Investigation,
vol. 6, pp. 13-15, cited in AFFA 2001a.
Larochelle, R., & Magar, R. 1997, ‘Evaluation of the presence of porcine reproductive and
respiratory syndrome virus in packaged pig meat using virus isolation and polymerase chain
reaction (PCR) method’, Vet Microbiol, vol. 58, pp. 1-8.
Magar, R., Robinson, Y., Dubuc, C., & Larochelle, R. 1995, ‘Evaluation of the persistence of
porcine reproductive respiratory syndrome virus in pig carcasses’, Vet Rec, vol. 137, pp. 559-561.
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Experts contacted:
Dr Ronald Magar
Health of Animals and Food Laboratory
Canadian Food Inspection Agency
St-Hyacinthe, Quebec
CANADA
P +1 450 773 7730
F +1 450 773 8152
[email protected]
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Western, Eastern and Venezuelan equine encephalomyelitis
Agent:
Family Togaviridae, genus alphavirus
Agent type:
Virus
Persistence and inactivation:
Equine encephalomyelitis viruses are transmitted by mosquitoes (Radostits et al 2000).
The level of viraemia produced in horses is very high (Radostits et al 2000), but has not been
reported to exceed 5 days in duration. The virus can be isolated from brain, although this has
proven unrewarding in some cases, or serum, which should be taken from pre-clinical febrile
horses (Walton 1992). The virus is present in saliva and nasal discharge.
References to the stability and inactivation of the virus were difficult to find – emphasis has
clearly been placed on trying to preserve the organism for diagnostic purposes rather than to
destroy it. Monath and Trent (1981) state that serum or tissue suspensions should be preserved at
-70oC. Alphaviruses will also survive at ambient temperatures for several days in blood dried on
paper discs. Equine encephalomyelitis viruses are extremely fragile and disappear from infected
tissues within a few hours after death (Radostits et al 2000).
Conclusions:
•
The equine encephalomyelitis viruses are fragile viruses that are not able to survive
for any length of time away from an animal host.
•
The disease is transmitted by mosquitoes.
•
Animal carcases and products pose a negligible transmission risk. No special
disposal measures are warranted.
References:
Monath, T.P., & Trent, D.W. 1981, ‘Togaviral diseases of domestic animals’, in Comparative
diagnosis of viral diseases, vol. IV, Vertebrate animal and related viruses. Part B – RNA viruses,
eds. Kurstak, E., & Kurstak, C., Academic Press, New York.
Radostits, O.M., Gay, C.G., Blood, D.C., & Hinchcliff, K.W. (eds) 2000, Veterinary medicine,
ninth edition, W. B. Saunders Company Ltd, London.
Walton, T.E. 1992, ‘Arboviral encephalomyelitides of livestock in the western hemisphere’, J Am
Vet Med Assoc, vol. 200, pp. 1385-1389.
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Other viruses
African swine fever
Agent:
Unclassified DNA virus like iridoviruses and poxviruses
Agent type:
Virus
Persistence and inactivation:
General characteristics: African swine fever virus is a very hardy agent. Stability has been
demonstrated in various strains for:
• six years at 5oC with no light
• eighteen months in serum at room temperature
• up to a month at 37oC
• 3.5 hours at 56oC (although serum can normally be safely sterilised after 30 minutes at
60oC).
ASFV is most stable at pH 4-10, but residual infectivity has been demonstrated at pH 3.1 after 22
hours, at pH 3.9 in serum after three days, and at pH 13.4 in serum after a week.
The virus is resistant to proteases (trypsin, pepsin), to nucleases and to putrefaction. It is very
sensitive to lipid solvents and detergents, as well as oxidising agents such as hypochlorite and
substituted phenols. Beta-propiolactone, acetyl-ethyleneimine and glycidaldehyde destroy
infectivity within one hour at 37oC, and formalin (0.5%) within about four days (Bengis 1997,
Farez and Morley 1997, Plowright and Parker 1967, Plowright et al 1994).
ASFV is present in nasal, oral, pharyngeal, conjunctival, genital, urinary and faecal secretions and
excretions. Transmission occurs by direct contact and via the environment (Plowright et al
1994).
Carcases and meat products: Kowalenko et al (1965) reported that ASFV persisted for 150 days
at 4oC and for 104 days at -4oC in skeletal muscle, and for six months in bone marrow at -4oC.
Persistence of ASFV in meat products has been comprehensively reviewed by Farez and Morley
(1997). The virus has been shown to survive for up to:
• 140 days in Iberian and white Serrano hams (Mebus et al 1993); Gregg (pers comm)
reported that in studies on salted and air-dried Serrano hams, starting at 4oC and working
up to room temperature, the virus survived about six months
• 399 days in Parma hams (McKercher et al 1987)
• 112 days in Iberian pork loins (Mebus et al 1993)
• 30 days in pepperoni and salami sausage (McKercher et al 1978).
Hams from acutely-infected animals should be safe if prepared using York or Parma procedures
and heated to 69oC for 3.5 hours or 70-75oC for 30 minutes. Smoked and spiced sausages or airdried hams require smoking to 32-49oC for 12 hours and drying for 25-30 days (Plowright et al
1994).
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Skins, hides and fibres: In a draft assessment, AFFA (2001) has concluded that unprocessed skins
and hides from susceptible animals in countries with ASFV pose a high risk. Given the hardy
nature of the virus, it is highly likely to be found on the skin of infected animals. Only fully
processed or fully tanned skins (including ‘wet whites’ and ‘wet blues’) are considered to pose no
quarantine threat, because the pH reached will inactivate the virus. Partially processed skins
(either limed or pickled) present a lower risk, but could not be relied on to be free of virus.
Semen/embryos: In a draft report, AQIS (2000) notes that there is little information on the
appearance and transmission of ASF in semen. Given the widespread secretion / excretion of the
virus from infected animals, infectivity of semen is likely, so the risk posed by semen is
considered moderate.
Faeces: Muller (1973) reported that ASFV may survive for 60-100 days in faeces, while
Eizenberger (unpublished, cited in Haas et al 1995) found that ASFV survived for at least 84 days
at 17oC and for 112 days at 4oC in slurry under simulated field conditions.
Conclusions:
•
The African swine fever virus is a persistent virus which can survive away from an
animal host for over a year under the right conditions.
•
Infection occurs mainly by the respiratory and oral routes, either directly with close
animal to animal contact or indirectly from the environment. Blood-sucking insects
can also spread the disease.
•
The ASF virus is highly contagious. Biosecurity of potentially infective material is
important.
•
Fresh carcases and meat from infected animals should be buried, burned or
rendered. If buried, the burial site must preclude scavenging by feral pigs.
Movement of carcases off-site for disposal should only be undertaken with a high
level of biological security during transportation.
•
Unprocessed skins that may be contaminated with ASF virus should be buried,
burned or disinfected prior to further processing. Fully processed skins are a
negligible disease risk.
•
Semen and embryos collected from infected animals should be destroyed.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 2000, Draft porcine semen risk analysis:
risk assessment and risk management options, March 2000, AQIS, Canberra.
Bengis, R.G. 1997, ‘Animal health risks associated with the transportation and utilisation of
wildlife products’, Rev sci tech Off int Epiz, vol. 16(1), pp. 104-110.
Blackwell, J.H. 1987, ‘Viruses in products of food animals’, Dairy and food sanitation, vol. 7, pp.
398-401.
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Haas, B., Ahl, R., Bohm, R., & Strauch, D. 1995, ‘Inactivation of viruses in liquid manure’, Rev
sci tech Off int Epiz, vol. 14(2), pp. 435-445.
Kowalenko, J., Sidorow, M., & Burba, L. 1965, ‘African swine fever and control measures’, Int
Zeitschr Landwirtsch Deutsche, vol. 1, pp. 47-52, cited in Blackwell 1987.
McKercher, P.D., Hess, W.R., & Hamdy, F. 1978, ‘Residual viruses in pork products’, Appl
Environ Microbiol, vol. 35, pp. 142-145, cited in Farez and Morley 1997.
McKercher, P.D., Yedloutschnig, R.J., Callis, J.J., Murphy, R., Panina, G.F., Civardi, A.,
Bugnetti, M., Fonn, E.H., Laddomada, A., Scarano, C., & Scatozza, F. 1987, ‘Survival of viruses
in ‘Prosciutto di Parma’ (Parma ham)’, Can Inst Food Sci Technol J, vol. 20(4), pp. 267-72, cited
in Farez and Morley 1997.
Mebus, C.A., Arias, M., Pineda, J.M., Tapiador, House, C., & Sanchez-Vizcaino, J.M. 1997,
‘Survival of several porcine viruses in different Spanish dry-cured meat products’, Food Chem,
vol. 59, pp. 555-559.
Muller, W. 1973, ‘Hygiene landwirtschaftlicher und kommunaler Abfallbeseitigungssystemen
[Hygiene of agricultural and municipal waste disposal systems]’, in Hohenheimer Arbeiten 69,
Ulmer, Stuttgart, Germany, cited in Haas et al 1995.
Plowright, W., & Parker, J. 1967, ‘The stability of African swine fever virus with particular
reference to heat and pH inactivation’, Arch Ges Virusforsch, vol. 21, pp. 383-402, cited in Farez
and Morley 1997.
Plowright, W., Thomson, G.R., & Neser, J.A 1994, ‘African swine fever’, in Infectious diseases
of livestock, Vol I, eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University
Press, Cape Town, pp. 568-599.
Experts contacted:
Dr Douglas Gregg
National Veterinary Service Laboratories, APHIS
Foreign Animal Disease Diagnostic Laboratory (Plum Island)
United States Department of Agriculture
PO Box 848
Greenport NY 11944
USA
P +1 613 323 3379
F +1 613 323 3366
[email protected]
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Aujeszky’s disease
Agent:
Family Herpesviridae, subfamily Alphaherpesvirinae
Agent type:
Virus
Persistence and inactivation:
General characteristics: Aujeszky’s disease virus is labile in the presence of heat, drying, and UV
light. It has a half-life of 7 hours at 37oC, but survives for long periods at <4oC (e.g. up to 46
days in contaminated straw and feeding troughs at -20oC)(Schoenbaum et al 1991). It is
destroyed by heating at 56oC within 30 minutes (Maré 1994). The virus is stable between pH 5-9
but is inactivated rapidly outside this range.
ADV is inactivated by most disinfectants, including sodium hypochlorite 0.5% (seconds),
phenolic derivatives 3% (10 minutes), and formaldehyde 0.6% (within one hour), and by lipid
solvents such as ethyl ether, acetone, chloroform, and alcohol. It is relatively resistant to sodium
hydroxide 0.8% and 1.6% (> 6 hours) (Pensaert & Klugge 1989).
ADV is spread via oral and nasal discharges, saliva and semen. Close contact facilitates spread
but airborne transmission over long distances may be possible. Ingestion of infected tissues and
foetuses may also give rise to infection in pigs, dogs, cats and wildlife (Maré 1994).
Carcases and meat products: Although it can be isolated from the tissues of infected pigs after
death (Heard 1980, Pensaert & Klugge 1989), ADV is not considered a high risk contaminant of
pig meat products. It does not appear in the OIE review by Farez & Morley (1997) of ‘potential
animal health hazards of pork and pork products’. MacDiarmid and Thompson (1997) noted that
sheep or goat meat infected with ADV would only pose a risk if fed uncooked to pigs, as dogs
and cats are dead-end hosts.
AFFA (2001a) has reviewed the literature on infectivity of ADV in pig meat, citing several
papers reporting the transmission of infection through the consumption of the carcases of infected
animals. ADV was recovered from the carcase muscle of clinically affected pigs after storage at
1-2oC for 72 hours (MacDiarmid 1991) but was inactivated in muscle, lymph node and bone
marrow from an artificially infected hindquarter after 35 days at -18oC (Durham et al 1980).
Wooley et al (1981) showed that ADV was inactivated after 72 hours in fermented edible food
wastes treated with Lactobacillus acidophilus, when temperatures were between 20-30oC. The
virus survived beyond 96 hours at temperatures between 5-10oC. Morrow et al (1995) concluded
that composting was effective at destroying ADV in pig carcases, provided that maximum
temperatures in the pile exceed 60oC.
Milk and milk products: This review did not uncover any specific reports on the spread of ADV
via milk, confirming the risk assessment of AQIS (1999a).
Skins, hides and fibres: This review did not uncover any specific reports on the persistence of
ADV on skins, hides or fibres. The risk of ADV being present for any period of time is likely to
be very small, given the susceptibility of the virus to drying and UV light. In a draft report,
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AFFA (2001b) has determined that “the mode of transmission of the disease is such that hides
and skins are unlikely to be a significant risk factor in the transmission [of Aujeszky’s disease]”.
Semen/embryos: ADV has been detected in the semen of boars and coital transmission is possible
(Maré 1994). In a draft report, AQIS (2000) have identified porcine semen as a high risk
quarantine threat for ADV.
AUSVETPLAN (DPIE 1996) notes that whilst ADV can attach to embryos (Bolin et al 1982),
trypsin treatment will remove the virus and transmission by embryo transfer under “natural
circumstances” is highly unlikely.
Faeces: A number of authors have studied the survival of ADV in pig and cattle slurry. Bøtner
(1991) reported findings reasonably consistent with previous studies. Inactivation increased with
temperature, requiring 15 weeks at 5oC, 2 weeks at 20oC, 5 hours at 35oC and 10 minutes at 55oC
(all at pH 7.3-7.9). The virus was inactivated more quickly in cattle than pig slurry (40 minutes
compared to 2.5 hours at 40oC) for unidentified reasons. Muller (1973) reported that ADV may
survive for 3-15 weeks.
Conclusions:
•
Aujeszky’s disease virus is a sensitive virus which, under normal conditions, does
not survive for more than a few days away from an animal host. It is readily
inactivated by drying, heat, UV light and commonly used disinfectants.
•
Infection principally occurs by the respiratory and oral routes, with close contact
required between infected and susceptible animals. Transmission can also occur
with infected semen.
•
The over-riding consideration with disposal of fresh carcases is that infective
material must not be consumed by pigs. Fresh carcases from infected animals
should be buried, burned or rendered. If buried, the burial site must preclude
scavenging.
•
The meat from clinically healthy animals on affected farms may be processed
normally.
•
Semen collected from infected animals should not be used.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001a, Generic import risk
analysis (IRA) for uncooked pig meat. Issues paper, January 2001, AFFA, Canberra.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001b, Import risk
analysis. Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999a, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999b, Importation of sausage casings into
Australia, import risk analysis, December 1999, AQIS, Canberra.
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AQIS (Australian Quarantine and Inspection Service) 2000, Draft porcine semen risk analysis:
risk assessment and risk management options, March 2000, AQIS, Canberra.
Bolin, S.R, Runnels, L.J., Sawyer, C.A., & Gustafson, D.P. 1982, ‘Experimental transmission of
pseudorabies virus in swine by embryo transfer’, Am J Vet Res, vol. 43, pp. 278-280, cited in
DPIE 1996.
Bøtner, A. 1991, ‘Survival of Aujeszky’s disease virus in slurry at various temperatures’, Vet
Microbiol, vol. 29, pp. 225-235.
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Aujeszky’s disease, DPIE, Canberra.
Durham, P.J.K., Gow, A., & Poole, W.S.H. 1980, ‘Survival of Aujeszky’s disease virus in frozen
pig meat’, Res Vet Sci, vol. 28, pp. 256-258, cited in AFFA 2001.
Farez, S., & Morley, R.S. 1997, ‘Potential animal health hazards of pork and pork products’, Rev
sci tech Off int Epiz, vol. 16(1), pp. 65-78.
Haas, B., Ahl, R., Bohm, R., & Strauch, D. 1995, ‘Inactivation of viruses in liquid manure’, Rev
sci tech Off int Epiz, vol. 14(2), pp. 435-445.
Heard, T.W. 1980, ‘Aujeszky’s disease virus in meat’, Vet Rec, vol. 107, p. 139.
MacDiarmid, S.C. 1991, ‘Importation into New Zealand of meat and meat products’, Regulatory
Authority , Ministry of Agriculture and Forestry, Wellington, New Zealand, cited in AFFA 2001.
MacDiarmid, S.C., & Thompson, E.J. 1997, ‘The potential risks to animal health from imported
sheep and goat meat’, Rev sci tech Off int Epiz, vol. 16(1), pp.45-56.
Maré, C.J. 1994, ‘Aujeszky’s disease’, in Infectious diseases of livestock, Vol II, eds. Coetzer,
J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 958-962.
Morrow, W.E.M., O’Quinn, P., Barker, J., Erickson, G., Post, K., & McCaw, M. 1995,
‘Composting a suitable technique for managing swine mortalities’, Swine Health Prod, vol. 3(6),
pp. 236-243.
Muller, W. 1973, ‘Hygiene landwirtschaftlicher und kommunaler Abfallbeseitigungssystemen
[Hygiene of agricultural and municipal waste disposal systems]’, in Hohenheimer Arbeiten 69,
Ulmer, Stuttgart, Germany, cited in Haas et al 1995.
Pensaert, M.B., & Klugge, J.P. 1989, ‘Pseudorabies virus (Aujeszky’s disease)’, in Virus
infections of vertebrates. Vol II Virus infections of porcines, ed. Pensaert, M.B., Elsevier Science
Publishers, Amsterdam, pp. 39-64.
Schoenbaum, M.A., Freund, J.D., & Beran, G.W. 1991, ‘Survival of pseudorabies virus in the
presence of selected diluents and fomites’, J Am Vet Med Ass, vol. 198, pp. 1393-1397, cited in
DPIE 1996.
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Wooley, R.E., Gilbert, J.P., Whitehead, W.K., Shotts, E.B., & Dobbins, C.N 1981, ‘Survival of
viruses in edible fermented waste materials’, Am J Vet Res, vol. 42, pp. 87-90.
Experts contacted:
Dr Sabrina Swenson
Head, Bovine and Porcine Viruses Section
Diagnostic Virology Laboratory
National Veterinary Services Laboratories
1800 Dayton Rd
Ames, Iowa, 50010
USA
P +1 515 663 7551
F +1 515 663 7348
[email protected]
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Borna disease
Agent:
Unclassified enveloped RNA virus
Agent type:
Virus
Persistence and inactivation:
General characteristics: There are many gaps in the knowledge about Borna disease and its
causative virus. Transmission is not well understood. BDV RNA has been detected in nasal
discharge, conjunctival fluid and saliva of horses (Richt et al 1993), but several attempts to
demonstrate infectivity in horse secretions have failed (Staeheli et al 2000). Infective virus has
been found in the urine of newborn infected rats (Morales et al 1988). Infection is thought to be
through ingestion or inhalation. BDV is sensitive to lipid solvents and UV light (Richt et al
1994).
According to Richt (pers comm), no work has been done on persistence of BDV in carcases or
animal products. No information was discovered in the literature review.
Carcases and meat products: Igata-Yi et al (1996), investigating a proposed association between
BDV and psychiatric disorders in Japanese people, were unable to find a link between the
consumption of raw horse meat and the presence of BDV. AQIS (1999) considered that the
transmission of Borna disease by meat or meat products appeared unlikely.
Skins, hides and fibres: No information on this topic was discovered during the conduct of this
review. In a draft assessment, AFFA (2001) has determined that the likelihood of transmission of
Borna disease via skins or hides is negligible.
Semen/embryos: No information on this topic was discovered during the conduct of this review.
Faeces: Muller (1973), using artificial inoculation, reported that Borna disease virus may survive
for 22 days in faeces. No other information on this topic was found.
Conclusions:
•
Little is known about the persistence of Borna disease virus in either the
environment or in animal products, and the epidemiology of the disease is poorly
understood.
•
The risk of disease transmission with animal products appears low.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999, Importation of sausage casings into
Australia, import risk analysis, December 1999, AQIS, Canberra.
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Haas, B., Ahl, R., Bohm, R., & Strauch, D. 1995, ‘Inactivation of viruses in liquid manure’, Rev
sci tech Off int Epiz, vol. 14(2), pp. 435-445.
Igata-Yi, R., Yamaguchi, K., Takemoto, S., Yamasaki, H., Matsuoaka, M., & Miyakawa, T. 1996,
‘Borna disease virus and the consumption of raw horse meat’, Nature Medicine, vol. 2, pp. 948949.
Morales, J.A., Herzog, S., Kompter, C., Frese, K., & Rott, R. 1983, ‘Axonal transport of Borna
disease virus along olfactory pathways in spontaneously and experimentally infected rats’, Med
Micro Immunol, vol.177, pp. 51-68, cited in Staeheli et al 2000.
Muller, W. 1973, ‘Hygiene landwirtschaftlicher und kommunaler Abfallbeseitigungssystemen
[Hygiene of agricultural and municipal waste disposal systems]’, in Hohenheimer Arbeiten 69,
Ulmer, Stuttgart, Germany, cited in Haas et al 1995.
Richt, J.A., Herzog, S., Haberzetti, K., & Rott, R. 1993, ‘Demonstration of Borna disease virusspecific RNA in secretions of naturally infected horses by the polymerase chain reaction’, Med
Micro Inmmunol, vol. 182, pp. 293-304, cited in Staeheli et al 2000.
Richt, J.A., Herzog, S., Schmeel, A., Frese, K., Clements, J.E., and Rott, R. 1994, ‘Current
knowledge about Borna disease and its causative virus’, in Equine infectious diseases VII,
Proceedings of the Seventh International Conference, Tokyo, eds. Nakajima, H., & Plowright, W.,
R&W Publications (Newmarket) Ltd, UK, pp. 55-60.
Rott, R., & Herzog, S. 1994, ‘Borna disease’, in Infectious diseases of livestock, Vol II, eds.
Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 978981.
Staeheli, P., Sauder, C., Hausmann, J., Ehrensperger, F., & Schwemmle, M. 2000, ‘Epidemiology
of Borna disease virus’, J Gen Virol, vol. 81, pp. 2123-2135.
Experts contacted:
Dr Juergen Richt
Veterinary Medical Officer
National Animal Disease Center – ARS – USDA
2300 Dayton Avenue
Ames, IA 50100
USA
P +1 515 663 7366
F +1 515 663 7458
[email protected]
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Classical swine fever
Agent:
Family Flaviviridae, genus pestivirus
Agent type:
Virus
Persistence and inactivation:
General characteristics: The survival and inactivation of classical swine fever virus has been
comprehensively reviewed. It can be described as moderately sensitive in the environment, but
persistent under the right conditions (cool, moist, and protein-rich), where it may survive for up to
a few weeks. It is rapidly inactivated in the presence of UV light (Edwards 2000).
Over a number of studies CSFV has been shown to be stable at neutral to slightly alkaline values
(around pH 5-10). It is rapidly inactivated below pH 3 and above pH 10 (Edwards 2000).
Thermal stability of CSFV varies with the media used, the pH and the strain of virus (Depner et al
1992, Edwards 2000). The virus is relatively labile at higher temperatures. Harkness (1985)
reported no loss of titre in cell culture fluid after 180 days at 4oC, -30oC, or -80oC, but inactivated
the virus after 30 minutes at 56oC or 10 minutes at 60oC. Inactivation took one minute at 90oC,
two minutes at 80oC and five minutes at 70oC in a study by ur Rehman (1987). Other studies
have shown longer inactivation times – for example, where defibrinated blood was the medium
used. CSFV is susceptible to rapid changes in temperature such as thawing and refreezing (Farez
and Morley 1997).
CSFV is an enveloped virus and therefore susceptible to lipid solvents and detergents such as
ether, chloroform, and deoxycholate, as well as chlorine-based disinfectants, phenolics,
quaternary ammonium compounds, and aldehydes (Edwards 2000).
The principal means of transmission of CSF is from pig to pig, by direct contact or through
feeding of infected swill. Infection enters via the oral and intranasal routes and can occur via
abraded skin or needle. There is a high level of virus in blood and tissues and saliva. The virus is
also excreted in smaller quantities in urine, faeces, nasal and ocular discharges (Terpstra 1994).
In over 100 cases of CSF managed in Lower Saxony, Germany, between 1993 and 1996, no
outbreaks were connected to the transport of carcases from infected holdings to rendering plants.
However outbreaks occurring within 1km of infected holdings were linked with the movement of
rodents after stamping out commenced (Rassow, pers comm).
Carcases and meat products: Pork provides a favourable environment for CSFV. It will survive
in pork and processed pork products for months, when meat is chilled, or for years if frozen
(Terpstra 1994).
CSFV has been reported to survive longer than four years in frozen pork (Edgar et al 1946), and
for up to 85 days in chilled fresh pork. Survival in food waste at room temperature may only be
several days (Edwards 2000).
Farez and Morley (1997) have thoroughly reviewed survival times of CSFV in various meat
products, for example:
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•
•
•
•
•
252 days in Iberian hams
140 days in Iberian shoulder and White Serrano hams
126 days in Iberian loins (Mebus et al 1997)
one month in the meat, and two months in the bone marrow of salt-cured pork
147 days in intestinal casings processed in water at 42.2oC for 30 minutes (Farez and
Morley 1997).
Traditional hams with long curing times should be safe (Edwards 2000). CSFV is inactivated in
meat by cooking for 30 minutes at 65oC, 15 minutes at 69oC, or one minute at 71oC (Farez and
Morley 1997 and Edwards 2000, citing several authors). Heating for 30 minutes at 62.5oC failed
to inactivate the virus so temperature control is critical (Edwards 2000).
Skins, hides and fibres: In a draft assessment, AFFA (2001b) has concluded that unprocessed
skins and hides from susceptible animals in countries with CSFV pose a high risk. The virus is
highly likely to be found on the skin of infected animals. Only fully processed or fully tanned
skins (including ‘wet whites’ and ‘wet blues’) are considered to pose no quarantine threat,
because the pH reached will inactivate the virus. Partially processed skins (either limed or
pickled) present a lower risk, but could not be relied on to be free of virus because of its stability
over a wide range of pH.
Semen/embryos: AQIS (2000) note that while there are reports of CSFV in semen in the
literature, they all seem to originate with a personal communication cited in Thacker et al (1984).
The virus was apparently isolated from an experimentally-infected boar and was shown to be
transmitted to a female. The virus was maintained by freezing.
AQIS (2000) has concluded that excretion of the virus in semen would not be surprising and that
the risk of transmission by artificial insemination should therefore be considered moderate.
Faeces: Bøtner (1990, cited in Haas et al 1995) found that time to inactivation for CSFV in slurry
varied between > 6 weeks at 5oC to instantaneous at 50oC (detection limit 0.7 log10 TCID50/50L).
Eizenberger (unpublished, cited in Haas et al 1995) found CSFV surviving for at least 70 days at
17oC and for 84 days at 4oC in slurry under simulated field conditions. Have (1984) reported loss
of infectivity after about 15 days in liquid slurry.
Terpstra (1994) states that CSFV appears to be inactivated within a few days in manure.
Conclusions:
•
The classical swine fever virus is a sensitive virus which, under normal conditions,
does not survive for more than a few days away from an animal host or animal
products. However, the virus can survive for a few weeks in a cool, moist proteinrich environment. It can survive for months in chilled pig meat and for years in
frozen pig meat.
•
Infection principally occurs by the respiratory and oral routes, with transmission by
direct contact or swill feeding.
•
All animal products potentially contaminated with the virus should be regarded as a
transmission risk.
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•
The over-riding consideration with disposal of fresh carcases is that infective
material must not be consumed by pigs.
•
On-site burning is the preferred method for disposal of carcases and animal
products.
•
Burial, burning and rendering off-site are less desirable options, which should only
be undertaken if transportation is biologically secure and scavenging by feral pigs
and rodents is precluded.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001a, Generic import risk
analysis (IRA) for uncooked pig meat. Issues paper, January 2001, AFFA, Canberra.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001b, Import risk
analysis. Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 2000, Draft porcine semen risk analysis:
risk assessment and risk management options, March 2000, AQIS, Canberra.
Bøtner, A. 1990, Modelstudier vederonde overlevelse af virus i gylle under traditionel opbevaring
og under udradning i biogasanlaeg. Research report, State Veterinaary Institute for Virus
Research, Lindholm, Denmark.
Depner, K., Bauer, T., & Liess, B. 1992, ‘Thermal and pH stability of pestiviruses’, Rev sci tech
Off int Epiz, vol. 11, pp. 885-893.
Edwards, S. 2000, ‘Survival and inactivation of classical swine fever virus’, Vet Microbiol, vol.
73, pp. 175-181.
Edgar, G., Hart, L., & Hayston, J.T. 1949, ‘Studies on the viability of the virus of swine fever’, in
Proceedings of the 14th International Veterinary Congress, London, pp. 387-391, cited in
Edwards 2000.
Farez, S., & Morley, R.S. 1997, ‘Potential animal health hazards of pork and pork products’, Rev
sci tech Off int Epiz, vol. 16, pp. 65-78.
Haas, B., Ahl, R., Bohm, R., & Strauch, D. 1995, ‘Inactivation of viruses in liquid manure’, Rev
sci tech Off int Epiz, vol. 14(2), pp. 435-445.
Harkness, J.W. 1985, ‘Classical swine fever and its diagnosis: A current view’, Vet Rec, vol. 11,
pp. 288-293, cited in AFFA 2001a.
Have, P. 1984, ‘Survival of swine fever virus in liquid manure’, FAO/EEC Expert Consultation
on Classical Swine Fever, Rome, October 1984, cited in Edwards 2000.
Mebus, C.A., Arias, M., Pineda, J.M., Tapiador, House, C., & Sanchez-Vizcaino, J.M. 1997,
‘Survival of several porcine viruses in different Spanish dry-cured meat products’, Food Chem,
vol. 59, pp. 555-559.
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Terpstra, C. 1994, ‘Hog cholera’, in Infectious diseases of livestock, Vol I, eds. Coetzer, J.A.W.,
Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 654-657.
Thacker, B.J., Larsen, R.E., Joo, H.S., & Leman, A.D. 1984, ‘Swine diseases transmissible with
artificial insemination’, J Am Vet Med Ass, vol. 185, pp. 511-516, cited in AQIS 2000.
Ur Rehman, S. 1987, ‚Untersuchungen zum viruziden Effekt bei der Erhitzung von
Speiseabfallen fur die Schweinemast’, Tierarztl Umsch, vol. 42, pp. 892-897, cited in Edwards
2000.
Experts contacted:
Dr John Pasick
Canadian Food Inspection Agency
National Centre for Foreign Animal Disease
1015 Arlington St
Winnipeg
Manitoba R3E 3M4
CANADA
P +1 204 789 2001
F +1 204 789 2038
[email protected]
Dr Gundula Floegel-Niesmann (on behalf of Professor Volker Moennig)
Institute of Virology
EU Reference Laboratory for CSF
D-30559 Hannover
GERMANY
P +49 511 953 8850
F +49 511 953 8898
[email protected]
Dr Dietrich Rassow
Veterinary Service
Oldenburg, Lower Saxony
GERMANY
[email protected]
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Infectious bursal disease (hypervirulent form)
Agent:
Family Birnaviridae
Agent type:
Virus
Persistence and inactivation:
General characteristics: Infectious bursal disease virus is highly resistant to environmental
influences and difficult to inactivate. It is resistant to freezing and thawing and is stable at pH >2
(Lukert and Saif 1997). Susceptible chickens have succumbed to the disease when introduced to
contaminated premises several weeks after depopulation (Geering et al 1995). IBDV is resistant
to many chemical disinfectants, but susceptible to chloramine solution, formalin and
glutaraldehyde (McFerran 1993).
Mandeville et al (2000) studied the heat lability of five strains of IBDV, both in vitro (Dulbecco’s
Modified Eagle Medium with 2% foetal calf serum) and by artificial inoculation of chicken
products. The in vitro study showed one, two, and three log reductions after heating for one
minute at 65oC, 71oC and 100oC respectively, with the greatest reduction occurring between 65oC
and 77oC. A similar thermal inactivation curve was observed after heating at 71oC or 74oC for up
to six minutes. The five strains showed similar characteristics.
Other heating studies have shown:
• apparent neutralisation after 30 minutes at 70oC (Landgraf et al 1967)
• little loss of infectivity after five hours at 56oC (Benton et al 1967)
• little loss of infectivity after 90 minutes at 60oC (Cho and Edgar 1969)
• reduction in infectivity of bursal homogenate supernatant by one log after 18.8 minutes at
70oC, after 11.4 minutes at 75oC, and after three minutes at 80oC (Alexander and Chettle
1998).
On the basis of work conducted on behalf of AQIS on IBDV inactivation in chicken meat, fat,
skin and bursa supernatant, imported chicken meat is required to be cooked for 165 minutes at
74oC or for 125 minutes at 80oC (AFFA 2000).
IBDV is shed in the faeces for up to two weeks following infection, and is transmitted through
oral infection or inhalation (Geering et al 1995, van den Berg 2000). Spread occurs with direct
contact or via fomites. Vertical transmission has not been demonstrated (Geering et al 1995).
There are two viraemic stages, the second leading to replication of the virus in several organs,
disease and often death (van den Berg 2000).
Carcases and meat products: No references were found on the persistence of IBDV in carcases.
According to AFFA (2001) and MAF (1999), IBDV has been found in the muscle of chickens
between 48 and 96 hours after infection in studies in Britain. It was also found in liver, kidney,
bursa, faeces and blood between 24 and 96 hours after infection. Mandeville et al (2000) note
that the quantity of IBDV present in chicken muscle during infection is not known, and that
studies using artificial inoculation must therefore be interpreted with caution.
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The only study to be carried out directly on chicken products has been that by Mandeville et al
(2000). Chicken products seeded with 105.5 TCID50/25µl of virus were heated to 71oC or 74oC
under simulated cooking conditions and quickly cooled after reaching target temperatures.
Infectivity was not destroyed. The authors noted that complete inactivation of IBDV would
require greater than six minutes at 71oC or 74oC. (This makes an interesting comparison with the
AQIS requirements that chicken meat be cooked for 165 minutes at 74oC or for 125 minutes at
80oC (AFFA 2000).)
Eggs and egg products: No references were found on the persistence of IBDV in or on eggs.
Vertical transmission is not thought to be a feature of the epidemiology of IBD, but
contamination of eggs with faeces is highly likely. On the basis of other data on environmental
survival, potential persistence on eggs is likely to be of the order of two to three months.
Other products: IBDV may survive for more than 60 days in poultry litter (Vindevogel et al
1976).
Conclusions:
•
Infectious bursal disease virus is a persistent virus that can survive in the
environment for months.
•
Infection occurs by the respiratory and oral routes, with transmission by direct
contact and fomites. The virus is highly contagious. Biosecurity of potentially
infective material is very important.
•
Burning, rendering and burial are the preferred methods for disposal of infective
material.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2000, The importation of
non-viable eggs and products containing egg. Technical issues paper, December 2000, AFFA,
Canberra.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Generic import risk
analysis (IRA) for uncooked chicken meat. Issues paper, July 2001, AFFA, Canberra.
Alexander, D.J., & Chettle, N.J. 1998, ‘Heat inactivation of serotype I infectious bursal disease
virus’, Avian pathol, vol. 27, pp. 97-99, cited in AFFA 2000 and Mandeville et al 2000.
Benton, W.J, Cover, M.S., Rosenberger, J.K., & Lake, R.S. 1967, ‘Physiochemical properties of
the infectious bursal agent (IRA)’, Avian Dis, vol. 11, pp. 2102-2109, cited in Mandeville et al
2000.
Cho, Y., & Edgar, S.A. 1969, ‘Characterisation of the infectious bursal agent’, Poultry Sci, vol.
48, pp. 560-565, cited in AFFA 2000 and Mandeville 2000.
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
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Landgraf, H., Vielitz, E., & Kirsch, R. 1967, ‘Occurrence of an infectious disease affecting the
bursa of Fabricius (Gumboro disease)’, Dtsch Tieraerztl Wochenschr, vol. 74, pp.6-10, cited in
Mandeville 2000.
Lukert, P.D., & Saif, Y.M. 1997, ‘Infectious bursal disease’, in Diseases of poultry, tenth edition,
ed. Calnek, B.W., Iowa State University Press, pp. 721-738, cited in MAF 1997.
MAF (Ministry of Agriculture and Forestry New Zealand) 1997, Import risk analysis: chicken
and chicken meat products; Bernard Matthews Foods Ltd turkey meat preparations from the
United Kingdom, Regulatory Authority, MAF, Wellington, New Zealand.
Mandeville III, W.F., Cook, F.K., & Jackwood, D.J. 2000, ‘Heat lability of five strains of
infectious bursal disease virus’, Poultry Science, vol. 79, pp. 838-842.
McFerran, J.B. 1993, ‘Infectious bursal disease’, in Virus infections of birds, eds. McFerran, J.B.,
& McNultry, M.S., Elsevier Science Publishers, Amsterdam, pp. 213-228.
Van den Berg, T.P. 2000, ‘Acute infectious bursal disease in poultry: a review’, Avian Pathol,
vol. 29, pp. 175-194.
Vindevogel, H., Gouffaux, M., Meulemans, G., Duchatel, J.P., & Halen, P. 1976, ‘Maladie de
gumboro: Distribution et persistance du virus chez le poussin inocule. Etudes sur la transmission
de la maladie’, Avian pathol, vol. 5, pp. 31-38, cited in Mandeville 2000.
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Transmissible gastroenteritis
Agent:
Family Coronaviridae
Agent type:
Virus
Persistence and inactivation:
General characteristics: Transmissible gastroenteritis virus is stable at -20oC, able to survive for
months at 4-5oC, but loses infectivity after 4 days at 37oC (Harada et al 1968, Pensaert 1989). It
survives down to pH 3 and is moderately sensitive to trypsin (Pensaert and Callebaut 1994).
TGEV can retain infectivity in the environment for up to three days (Geering et al 1995) or “no
longer than a few weeks” (Radostits et al 2000). The virus is photosensitive (Radostits et al
2000).
TGEV is sensitive to lipid solvents and detergents and to a wide range of disinfectants, including
sodium hypochlorite, sodium hydroxide, formaldehyde solution (including vapour), iodine,
phenolic and quaternary ammonium compounds (Brown 1981).
TGE is highly contagious. It is principally spread by the faecal-oral route. While several authors
note that transmission may take place via aerosols (e.g. Forman 1991), Pensaert and Callebaut
(1994) state that spread of the virus by this means is not known to occur. The virus replicates in
the respiratory tract and is present in aerosols, but in lower concentrations than in faeces. Spread
is by direct contact or indirect through the environment (Geering et al 1995, Pensaert and
Callebaut 1994). Virus has also been found in the milk of infected sows (Kemeny et al 1975).
Pigs are the only species affected, and although dogs, cats and foxes have been infected
experimentally, it is unlikely they play a role in the epidemiology (Pensaert and Callebaut 1994).
Carcases and meat products: TGEV has been found in a range of tissues. Cook et al (1991) cite
an early study in which TGE was transmitted by homogenates of kidney, spleen, liver, lungs,
brain, and gastrointestinal tract (Bay et al 1949), and another in which virus was isolated in nasal
and tracheal mucosa, oesophagus, lung, intestine, and lymph nodes (Harada et al 1969).
Forman (1991) succeeded in transmitting infection by feeding ground-up samples of muscle, bone
marrow and lymph node from recently infected pigs to one- and three-week-old piglets. The
tissues had been stored at -25oC for at least 30 days. The conditions were intended to mimic real
abattoir conditions. The author concluded that the risk of transmission by this means was low but
real.
In a similar study, Cook et al (1991) demonstrated transmission from apparently healthy animals
from a TGE-endemic area. TGE virus was found in tonsils from 4 of 500 pigs but not from
lymph nodes or muscle. The study showed the risk of introducing TGE through pig carcases
from TGE-endemic areas even when they have passed ante-mortem examination.
The acid stability of TGEV means it is not inactivated by rigor mortis or lactic fermentation of
certain meat products (Harada et al 1968, Leman et al 1986). Putrefaction is known to destroy
the virus (Leman et al 1986). Sausage casings are considered unlikely to transmit TGE because
they are salted and stored at room temperature, although no direct evidence could be found for
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this (AQIS 1999). References on the effect of cooking or curing on the TGE virus were not found
(see also AFFA 2001a), although AUSVETPLAN (DPIE 1996) states that TGEV will be
destroyed by cooking.
Skins, hides and fibres: No specific reports were found on this aspect of TGE. In a draft
assessment, AFFA (2001b) have concluded that skins and hides pose an insignificant risk in the
transmission of TGE.
Semen/embryos: There are no reports of TGE virus in the semen of boars, or spread of the disease
via artificial insemination. As the virus is excreted in the faeces, there is some risk of semen
contamination from this source, but this risk is considered to be low (AQIS 2000).
Faeces: TGEV is excreted in the faeces for around 14 days (Pensaert et al 1970). Faeces
containing the virus was no longer infective after 10 days at 21oC (Young et al 1955). Faeces
containing 105 TGEV was inactivated within 6 hours when exposed to direct sunlight (DPIE
1996).
Conclusions:
•
The TGE virus is a sensitive virus which, under normal conditions, can survive in
the environment for only a few days. It is inactivated by commonly used
disinfectants.
•
TGE is highly contagious. The disease has an oral-faecal cycle.
•
Fresh carcases must be disposed of in a manner that precludes scavenging by pigs.
•
Provided there is no swill feeding of pigs, meat and skins pose a negligible
transmission risk. No special disposal precautions are warranted.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001a, Generic import risk
analysis (IRA) for uncooked pig meat. Issues paper, January 2001, AFFA, Canberra.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001b, Import risk
analysis. Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999, Importation of sausage casings into
Australia, import risk analysis, December 1999, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 2000, Draft porcine semen risk analysis:
risk assessment and risk management options, March 2000, AQIS, Canberra.
Bay, W.W., Hutchings, L.M., Doyle, L.P., & Bunnell, D.E. 1949, J Am Vet Med Assoc, vol. 115,
p. 245, cited in Cook et al 1991.
Brown, T.T. 1981, ‘Laboratory evaluation of selected disinfectants as virucidal agents against
porcine parvovirus, pseudorabies virus and transmissible gastroenteritis virus’, Am J Vet Res, vol.
36, pp. 267-271. cited in AQIS 1999.
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Cook, D.R., Hill, H.T., & Taylor, J.D. 1991, ‘Oral transmission of transmissible gastroenteritis
virus by muscle and lymph node from slaughtered pigs’, Aust Vet J, vol. 68, pp. 68-70.
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Transmissible Gastroenteritis, DPIE,
Canberra.
Forman, A.J. 1991, ‘Infection of pigs with transmissible gastroenteritis virus from contaminated
carcases’, Aust Vet J, vol. 68, pp. 25-27.
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
Harada, K., Kaji, T., Kumagai, T., & Sasahara, J. 1968, ‘Studies on transmissible gastroenteritis
in pigs. IV. Physiochemical and biological properties of TGE virus’, Natl Inst Anim Health Q
(Tokyo), vol. 8, pp. 140-147, cited by Cook et al 1991 and AQIS 1999.
Harada, K., Furuuchi, S., Kumagai, T., & Sasahara, J. 1969, Natl Inst Anim Health Q (Tokyo),
vol. 9, p. 185, cited by Cook et al 1991.
Kemeny, L.J., Wiltsey, V.L., & Riley, J.L. 1975, ‘Upper respiratory infection of lactating sows
with transmissible gastroenteritis virus following contact exposure to infected pigs’, Cornell Vet.,
vol. 65, pp. 352-362, cited in Forman 1991 and DPIE 1996.
Leman, A.D., Straw, B., Glock, R.D., Mengeling, W.L., Penny, R.H.C., & Scholl, E. 1986, in
Diseases of Swine, sixth edition, Iowa State University Press, Iowa, USA, cited by AFFA 2001a.
Pensaert, M.B. 1989, ‘Porcine epidemic diarrhoea virus’, in Virus infections of vertebrates. Vol 2
Virus infections of porcines, ed. Pensaert, M.B., Elsevier Science Publishers, Amsterdam, pp.
167-176.
Penseart, M.B., & Callebaut, P. 1994, ‘Transmissible gastroenteritis’, in Infectious diseases of
livestock, Vol II, eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press,
Cape Town, pp. 866-869.
Pensaert, M.B, Haelterman, E.O., & Burnstein, T. 1970, ‘Transmissible gastroenteritis of swine:
Virus-intestinal cell interactions. I. Immunofluorescence, histopathology and virus production in
the small intestine through the course of the infection’, Arch Gesamte Virusforsch, vol. 31, pp.
321-334, cited in Forman 1991.
Radostits, O.M., Gay, C.G., Blood, D.C., & Hinchcliff, K.W. (eds) 2000, Veterinary medicine,
ninth edition, W. B. Saunders Company Ltd, London.
Young, G.A., Hintz, R.W., & Underdahl, N.R. 1955, ‘Some characteristics of transmissible
gastroenteritis in disease free antibody-devoid pigs’, Am J Vet Res, vol. 16, pp. 529-535, cited in
DPIE 1996.
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Vesicular exanthema
Agent:
Family Caliciviridae
Agent type:
Virus
Persistence and inactivation:
General characteristics: Vesicular exanthema is relatively resistant to environmental influences. It
may survive for up to 2 years at refrigerator temperatures, and up to 6 weeks at room temperature
(Madin 1989). It is inactivated after 60 minutes at 62oC or 30 minutes at 64oC, and outside pH
range 3 to 9 (MacDiarmid 1991).
According to Madin (1989), farms that are heavily contaminated must be regarded as infectious
for several months unless there is vigorous disinfection. However, Bankowski (1965) reported
the case of a farm on which there was no evidence of infectivity seven days after depopulation.
VESV is susceptible to most common disinfectants, but not detergents (DPIE 1996). Madin
(1989) specifically nominates VESV sensitivity to 2% sodium hydroxide, 0.1% sodium
hypochlorite, and 2% citric acid. Manure, fat and other organic matter is protective and must be
removed prior to disinfection (DPIE 1996).
VESV is excreted in the saliva and faeces of infected pigs, from about 12 hours prior to clinical
signs and for 1-5 days after that (Wilder and Dardiri 1978). Vesicles containing virus also
rupture. Transmission can be by direct, but not indirect, contact between pigs, and via the
ingestion of untreated swill, but whether the infection establishes via the oral route or via skin is
unknown. Infection has been established through scarified skin. VESV is found in a wide range
of tissues of infected pigs (Bankowski 1965, Thomson 1994).
Carcases and meat products: Viral particles can be identified in the muscle of slaughtered pigs
during the viraemic period (Wilder and Dardiri 1978). Patterson and Songer (1954) showed that
feeding infected muscle tissue, lymph node, heart muscle, spleen, lung, kidney, blood and crushed
bone to susceptible pigs can cause infection. The US was apparently unable to eradicate VES
until cooking garbage before feeding to swine became mandatory (Madin 1989).
Mott et al (1953) demonstrated survival of VESV in meat scraps of up to 4 weeks at 7oC, and at
for 18 weeks at -70oC.
Madin (1989, citing Traum and White unpublished 1941) noted that cooking meat at 184oF
(84.5oC) under 10lb (4.5kg) pressure did not destroy infectivity. However, MacDiarmid (1991)
reported that meat would be safe after 2-3 minutes at 80-100oC or 25 minutes at >70oC.
Radostits et al (2000) state that eradication should involve the slaughter of infected animals, but
that carcases can be salvaged for human consumption provided they are treated to inactivate the
virus.
Skins, hides and fibres: No specific reports were found on this aspect of VES (see also AFFA
2001b).
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Semen/embryos: Vesicular exanthema has been reported in boar semen (Cartwright and Huck
1967), but it is thought unlikely that the disease could be spread by artificial insemination (Hare
1985). Transmission via semen is not noted in any of the reviews examined during this study
(Geering 1995, Madin 1989, Radostits 2000, Thomson 1994). AQIS (2000) has concluded that
the risk of transmission of VES via artificial insemination is low.
Faeces: Virus is found in the faeces of infected pigs (Wilder and Dardiri 1978). No specific
reports were found on its persistence or inactivation in faeces.
Conclusions:
•
The vesicular exanthema virus is a persistent virus which can survive for several
weeks away from an animal host or animal products. The virus can survive for
months in chilled pig meat and for years in frozen pig meat.
•
Infection occurs mainly by the oral route, with transmission by direct contact or
swill feeding.
•
All animal products potentially contaminated with the virus should be regarded as a
transmission risk.
•
The over-riding consideration with disposal of carcases and animal products is that
infective material must not be consumed by pigs. Burial, burning and rendering are
all suitable methods of disposal. If infective material is buried, the site must
preclude scavenging by feral pigs.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001a, Generic import risk
analysis (IRA) for uncooked pig meat. Issues paper, January 2001, AFFA, Canberra.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001b, Import risk
analysis. Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 2000, Draft porcine semen risk analysis:
risk assessment and risk management options, March 2000, AQIS, Canberra.
Bankowski, R.A. 1965, ‘Vesicular exanthema’, Adv Vet Sci, vol. 10, pp. 23-64, cited in DPIE
1996 and Thomson 1994.
Cartwright, S.F., & Huck, R.A. 1967, ‘Viruses isolated in association with herd infertility,
abortions and stillbirths in pigs’, Vet Rec, vol. 81, pp. 196-197, cited in AQIS 2000.
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Vesicular Exanthema, DPIE, Canberra.
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
Hare, W.C.D. 1985, ‘Diseases transmissable by semen and embryo transfer techniques’,
Technical Series 4, Office International des Epizooties, Paris, cited in AQIS 2000.
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MacDiarmid, S. C. 1991, The importation into New Zealand of meat and meat products, Ministry
of Agriculture and Fisheries (MAF), New Zealand, cited in DPIE 1996.
Madin, S.H. 1989, ‘Vesicular exanthema virus’, in Virus infections of vertebrates. Vol 2 Virus
infections of porcines, ed. Pensaert, M.B., Elsevier Science Publishers, Amsterdam, pp. 267-271.
Mott, L.O., Patterson, W.C., Songer, J.R. & Hopkins, S.R. 1953, ‘Experimental infections with
vesicular exanthema’, in Proceedings of the 58th annual meeting of the US Livestock Sanitary
Association, pp. 334-340, cited in AFFA 2001a.
Patterson, W.C., & Songer, J.R. 1954, ‘Experimental infection with vesicular exanthema of swine
III: Viraemia studies in swine and their relationship to vesiculation’, in Proceedings of the 58th
annual meeting of the US Livestock Sanitary Association, pp. 294-301, cited in AFFA 2001a.
Radostits, O.M., Gay, C.G., Blood, D.C., & Hinchcliff, K.W. (eds) 2000, Veterinary medicine,
ninth edition, W. B. Saunders Company Ltd, London.
Thomson, G.R. 1994, ‘Vesicular exanthema’, in Infectious diseases of livestock, Vol II, eds.
Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 774775.
Wilder, F.H., & Dardiri, A.H. 1978, ‘San Miguel sea lion virus fed to mink and pigs’, Can J
Comp Med, vol. 42, pp. 200-204, cited in AFFA 2001a.
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Prions
Bovine spongiform encephalopathy and scrapie
Agent:
BSE and scrapie agents
Agent type:
Prion
Persistence and inactivation:
General characteristics: BSE agent has so far only been found only in CNS tissue, distal ileum
and bone marrow. However the much wider distribution of agent in kuru, Creuzfeld-Jakob
disease and scrapie suggests that all tissues should be treated with caution (Brown 1998).
The concentration of infective agent in TSEs is extremely high in naturally occurring disease.
The normal, conservative estimate used for oral infective dose (ID50) of BSE is 0.1g infected CNS
tissue for cattle, 1g for humans. Bovine brain and spinal cord weigh approximately 750g, so
assuming 1000g for risk assessment purposes, there are 10,000 infective oral doses of BSE per
cow (MAFF 2000).
Transmissible spongiform encephalopathy agents are extremely difficult to inactivate. In Iceland,
scrapie has been contracted by sheep grazing pastures that had lain unused for three years after
having been grazed by scrapie-infected sheep (Palsson 1979). Virtually all conventional
disinfection methods have failed to reliably and completely inactivate TSE agents (Taylor 2000),
including:
• irradiation and UV light
• dry heat, even to 360oC for one hour (drying of tissue is known to enhance its
thermostability)
• autoclaving, although gravity-displacement autoclaving at 132oC for 4.5 hours has been
recommended, as has porous-load autoclaving at 134-138oC for 18 minutes
• acids and bases, including hydrochloric acid (pH 0.1) for one hour and 2M sodium
hydroxide for two hours, although combinations of sodium hydroxide and heat have been
effective (see below)
• alkylating agents, including formalin and glutaraldehyde
• detergents, although boiling in SDS has been reported as effective for fluids
• halogens, except strong sodium hypochlorite (20,000ppm available chlorine for at least
one hour)
• organic solvents, including acetone, chloroform, and phenolics
• oxidising agents, including hydrogen peroxide and peracetic acid
• salts, including potassium permanganate
• chaotropes, including urea
• proteolytic enzymes, including trypsin, although pronase and proteinase K may reduce
titre significantly over prolonged exposure periods.
Taylor (2000) concluded that disinfection procedures that efficiently and/or rapidly fix proteins
(alcohols, aldehydes, rapid steam heating) enhance the thermostability of TSE agents. The only
completely effective methods were strong (20,000ppm) sodium hypochlorite solutions, applied
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for one hour, boiling in 1M sodium hydroxide for at least one minute, or gravity-displacement
autoclaving in the presence of sodium hydroxide (e.g. 121oC for 30-60 minutes plus 1M or 2M
NaOH).
Brown (1998) stated that, in regard to BSE-infected materials, “a fairly obvious recommendation,
based on the science, would be that all material that is actually or potentially contaminated with
BSE, whether whole carcases, rendered solids, or waste effluents, should be exposed to lye
[sodium hydroxide] and thoroughly incinerated under strictly inspected conditions. Another is
that the residue is buried in landfills to a depth that would minimise any subsequent animal or
human exposure, in areas that would not intersect with any potable water-table source”.
Carcases and meat products: Studies have detected residual scrapie infection in the packaging and
adjacent soil around scrapie-infected hamster brains buried for three years. Temperatures at the
site ranged from -20oC to 40oC (average 3oC to 25oC), with rainfall of 1000mm. Infectivity was
reduced by 98-99%. There was little leaching from the initial site, with no infectivity detected
more than 8cm from the burial site (Brown and Gajdusek 1991).
In a series of reports for the UK’s Environment Agency, Det Norska Veritas Ltd (Environment
Agency 1997a-e) estimated the risks posed to human health via environmental pathways, and of
various options for disposal of material potentially contaminated with BSE. The assumptions
may now be out of date, and the data used specific to the sites modelled. However the approach
taken provides a useful framework for comparing various disposal options.
In a risk assessment of landfills containing BSE-infected carcases from prior to 1991,
Environment Agency (1997d) concluded that “risk estimates for contamination of water supply
with BSE infectivity are all well below any level that would be considered to be of any
significance”. The calculated risk for the most exposed individual ranged from one in 10,000
million years to one in one million years3. The greatest risk came from leachate contaminating
water supplies. A favourable factor is that prions tend not to move with leachate, instead tending
to stick to other proteinaceous material or solids (consistent with Brown and Gajdusek 1991).
The risk assessment on disposal of BSE-infected carcases in incinerators estimated a maximum
risk of individual human infection of 1 in 1000 million (Environment Agency 1997c). Residual
infectivity after incineration was assumed to be 0.002%.
Risk assessments have also been conducted for treated waste water from plants rendering cattle
from the Over Thirty Months Scheme (Environment Agency 1997a, MAFF 2000) and for burning
rendered products from the OTMS in power stations (Environment Agency 1997b). An overview
of public health risks from BSE via environmental pathways in the UK has also been published
(Environment Agency 1997e).
The policy in the UK is to dispose of BSE-infected tissues by incineration. Landfill has not been
used for disposal since 1991 (DEFRA 2001). The UK currently has about 200,000 tons of ash
left from incinerated carcases from the foot-and-mouth eradication campaign. The pyres reached
high temperatures (evidenced by the fragmentation of bones) but there is concern that the animals
3
The risks quoted by the Environment Agency have been put into perspective by Gunn (2001), who noted
that the probability of dying from cancer was one in 300, and from being involved in a railway accident one
in 500,000.
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slaughtered could potentially include pre-clinical cases of BSE. The ash will therefore be
incinerated as a precaution (Jeffrey pers comm).
Milk and milk products: No reports were found of scrapie or BSE agents being present in milk.
AQIS (1999) has accepted the conclusions of the OIE that scrapie and BSE are not transmitted by
milk.
Skins, hides and fibres: No reports were found of scrapie or BSE agents contaminating skins,
hides or fibres. The agents would not be expected to be present on these products. In a draft
assessment, AFFA (2001) has concluded that skins, hides and fibres pose no quarantine hazard
for the introduction of scrapie or BSE.
Semen/embryos: The risk of transmission of BSE and scrapie via semen and embryos has been
reviewed by Wrathall (1997). A number of studies have been unable to detect scrapie agent in
ram semen or to demonstrate venereal transmission from infected rams (Palmer 1959, Foote
unpublished cited in Wrathall 1997), but these studies had shortcomings and further work is
needed. There appears to be no published work on scrapie in goat semen. Transmission of BSE
via semen has not been observed in either experimental conditions or by analysis of mating
records in Britain, and the risk is thought to be small or non-existent (Wrathall 1997).
The literature on possible transmission of TSEs via embryos is more complicated. Again, there
are shortcomings in published studies on scrapie in sheep and goats from groups in the USA and
Scotland (see for example Foote et al 1993, Foster et al 1996), and it is still not possible to say
whether scrapie is transmitted via embryos. A study by Foster et al (1999) failed to transmit
experimental BSE in goats via embryos. A large trial on transmission of BSE by embryo transfer
in cows in the UK had shown no transmission of the agent more than five years after the first
transfers (Wrathall 1997).
Faeces: No reports were found of scrapie or BSE agents being present in faeces.
Conclusions:
•
The TSE diseases are caused by prions, which are extremely hardy. Prions can
suvive in the environment for many years and are resistant to all conventional
methods of disinfection.
•
Infection occurs by the oral route, with an exceedingly small infective dose.
•
Carcasses should be burned completely and residual ash collected, mixed with
agricultural lime and deep buried at a suitable site which will need to be monitored
and scavengers excluded.
•
When burning is not practical, carcasses and other materials not able to be
decontaminated, should be deep buried with caustic materials at a suitable site
which will need to be monitored and scavengers excluded.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
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AQIS (Australian Quarantine and Inspection Service) 1999, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 2000, Import risk analysis report on the
revision of import policy related to scrapie, final report, August 2000, AQIS, Canberra.
Brown, P. 1998, ‘BSE: the final resting place’, Lancet, vol. 351, pp. 1146-1147.
Brown, P., & Gajdusek, D.C. 1991, ‘Survival of scrapie after 3 years’ interment’, Lancet, vol.
337, pp. 219-220.
Brown, P., Liberski, P.P., Wolff, A., & Gajdusek, D.C. 1990, ‘Resistance of scrapie infectivity to
steam autoclaving after formaldehyde fixation and limited survival after ashing at 360oC:
practical and theoretical implications’, J Infect Dis, vol. 161, pp. 467-472.
DEFRA (Department of Environment, Food and Rural Affairs UK), 2001, web site
www.defra.gov.uk.
Environment Agency (UK) 1997a, Thruxted Mill rendering plant – risk assessment of waste
water disposal options, report by Det Norska Veritas Ltd, London.
Environment Agency (UK) 1997b, Risks from burning rendered products from the over thirty
month scheme in power stations, report by Det Norska Veritas Ltd, London.
Environment Agency (UK) 1997c, Risks from disposing of BSE infected cattle in animal carcase
incinerators, report by Det Norska Veritas Ltd, London.
Environment Agency (UK) 1997d, Assessment of risk from BSE carcases in landfills, report by
Det Norska Veritas Ltd, London.
Environment Agency (UK) 1997e, Overview of risks from BSE via environmental pathways,
report by Det Norska Veritas Ltd, London.
Foote, W.C., Clark, W., Maciulis, A., Call, J.W., Hourrigan, J., Evans, R.C., Marshall, M.R., &
de Camp, M. 1993, ‘Prevention of scrapie transmission in sheep, using embryo transfer’, Am J vet
Res, vol. 54, pp. 1863-1868, cited in Wrathall 1997.
Foster, J.D., Hunter, N., Williams, A., Mylne, M.J.A., H. McKelvey, W.A.C., Hope, J., Fraser,
H., & Bostock, C. 1996, ‘Observations on the transmission of scrapie in experiments using
embryo transfer’, Vet Rec, vol. 138, pp. 559-562, cited in Wrathall 1997.
Foster, J.D., McKelvey, W.A.C., Fraser, H., Chong, A., Ross, A., Parnham, D., Goldmann, W.,
and Hunter N. 1999, ‘Experimentally induced bovine spongiform encephalopathy did not transmit
via goat embryos’, J gen Virol, vol. 80, pp. 517-524, cited in AQIS 2000.
Gale, P., & Stanfield, G. 2001, ‘Towards a quantitative risk assessment for BSE in sewage
sludge’, J Appl Microbiol, vol. 91, pp. 563-569.
Gunn, M. 2001, ‘Observations on disposal of BSE-infected carcasses’, Ir Vet J, vol. 54, pp. 192193.
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MAFF (Ministry of Agriculture, Fisheries and Food) 2000, Risk assessment for the disposal of
treated rendering plant ruminant condensate to agricultural land, report by Gale, P., Dee, A., &
King, P., sourced from DEFRA 2001.
Palmer, A.C 1959, ‘Attempt to transmit scrapie by injection of semen from an affected ram’, Vet
Rec, vol. 71, p. 664, cited in Wrathall 1997.
Palson, P.A. 1979, ‘Rida (scrapie) in Iceland and its epidemiology’, in Slow transmissible
diseases of the nervous system, vol I, eds. S. Prusiner & W. Hadlow, Academic Press, New York,
pp. 357-366, cited in Brown 1998.
Taylor, D.M. 2000, ‘Inactivation of transmissible degenerative encephalopathy agents: a review’,
Vet J, vol. 159, pp. 10-17.
Wrathall, A.E. 1997, ‘Risks of transmitting scrapie and bovine spongiform encephalopathy by
semen and embryos’, Rev sci tech Off int Epiz, vol. 16(1), pp. 240-264.
Experts contacted:
Dr Martin Jeffrey
Veterinary Laboratories Agency
Pentlands Science Park
Bush Loan, Penicuik
Midlothian EH26 0PZ
UK
P +44 131 445 6169
F +44 131 445 6166
[email protected]
Dr Robert Somerville
Neuropathogenesis Unit
Institute for Animal Health
West Mains Rd
Edinburgh EH9 3JF
UK
P +44 131 319 8235
F +44 131 668 3872
[email protected]
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Bacteria, mycoplasmas and fungi
Anthrax
Agent:
Bacillus anthracis
Agent type:
Bacterium
Persistence and inactivation:
General characteristics: Bacillus anthracis is an aerobic, spore-forming bacillus. It is normally
transmitted by the ingestion of spores, but infection can also take place via wounds and insect
bites. Infection through inhalation is thought to be rare (de Vos 1994), and windborne spread of
spores is minimal (Turnbull et al 1998).
Entry of B. anthracis to the body is followed by replication in the regional lymph nodes,
septicaemia, and invasion of all body tissues (Radostits et al 2000).
Spores are never found in the living body. Sporulation is a response to nutrient depletion from
drying of tissues and aerosolisation of fluids, with the opening of the carcase allowing ‘escape’ to
an aerobic environment (Dragon and Rennie 1995). Sporulation is inhibited by high partial
pressure of CO2 and temperatures below 20oC, and only takes place under appropriate conditions,
most notably when an infected carcase is opened. Germination of spores occurs at 20-40oC and at
relative humidity greater than 80% (de Vos 1994).
The vegetative (growing) form of B. anthracis is relatively labile. It is killed by 60oC dry heat for
30 minutes. Spores are much more robust, requiring 140oC dry heat for up to 3 hours for
inactivation (Buxton and Fraser 1977). Moist heat destroys spores at 100-115oC after 14.2
minutes (Bohm 1990).
Anthrax spores are generally resistant to alcohols, phenols, quaternary ammonium compounds,
ionic or non-ionic surfactants, acids and alkalis (Bengis 1997, de Vos 1994). They are
susceptible to 10% hot caustic soda, 4% formaldehyde, chlorine-containing disinfectants, 7%
hydrogen peroxide, and 2% glutaraldehyde (Whitford 1987).
De Vos (1994) cites several factors that impact on the survival of survival of anthrax spores in the
environment: initial numbers, topography, climate, and the presence of certain chemicals, other
microbes and plant material. In soils of high biological activity, survival may only be of the order
of 3-4 years (Whitford 1978). Under ideal conditions, spores may survive almost indefinitely in
the environment. Infective spores determined to be 200 years old have been found (de Vos
1994).
Carcases and meat products: All parts of the carcase of an anthrax victim may be infective. In the
unopened carcase, survival of the organism is short. Estimates include:
• no longer than 3 days at 25-30oC or higher (Stein 1947)
• two weeks in the skin, one week in the bone marrow (Minett 1950)
• up to 4 weeks at temperatures of 5-10oC (Whitford 1978).
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Opening of the carcase by scavengers or humans usually triggers sporulation before all of the
vegetative forms have been inactivated. The concept of survival of anthrax spores in carcases is
almost meaningless, as the spores may persist much longer than the carcase itself. A number of
factors – scavengers, water, wind – act to disseminate the spores into the environment (Dragon
and Rennie 1995).
Early identification and disposal of carcases prior to opening helps to minimise environmental
contamination. Carcases should be incinerated intact, or buried in a 2 metre deep grave, covered
with one part of chloride of lime (at least 25% active chlorine) to three parts soil (de Vos 1994).
The WHO (1994) has noted that burning is the most reliable method of destroying spores when it
is correctly done. No specific studies were found of the survival or inactivation of anthrax spores
in carcases after burning.
Bone meal contaminated with anthrax was found to be infective after 15 minutes of steam
treatment at 115oC degrees or 3 hours with dry heat at 140oC (De Kock et al 1940). WHO
guidelines for disinfection of bone meal (Whitford 1987) include several alternatives:
• steam sterilisation at 2.7 bar for 2 hours in a digester (less than 4 ton capacity)
• benzene vapour at 95-115oC for at least four hours, followed by 2 hours with live steam
at 5.4 bar (for broken bones)
• benzene vapour for eight hours at 95-115oC (for broken bones).
Heat sterilisation of bone meal for at least 3 hours should also be effective (Whitford 1978).
Milk and milk products: No specific statements were found in the literature about the excretion of
anthrax bacteria in milk. AQIS (1999) states that anthrax is not known to be transmitted in dairy
products.
Milk production decreases in lactating cows and the residual milk is either blood-stained or
yellow (de Vos 1994). Radostits et al (2000) stress the importance of preventing milk from
entering the human food chain. WHO guidelines (Whitford 1987) advise that milk should can be
considered decontaminated by adding chloride of lime (at least 25% active chlorine) at 1kg/20L
of milk for six hours.
Skins, hides and fibres: Numerous outbreaks of anthrax have been traced to infected hides and
associated products (de Vos 1994). In fact, de Vos (1994) states that hides from carcases of
infected animals should be regarded as permanently infected with B. anthracis.
Whitford (1987) gives details of the WHO guidelines for disinfection of infected materials. Wool
should be disinfected with formaldehyde, and hides with a mixture of 2.5% hydrochloric acid and
15% salt. Hair may be boiled, autoclaved at 120oC for 20 minutes, dry heated at 95oC for 24
hours, or steamed for six hours.
AFFA (2001) lists methyl bromide, formaldehyde (4-5%), glutaraldehyde, hydrogen peroxide,
peracetic acid and gamma radiation (4 MRad) as being capable of sterilising anthrax spores.
Later WHO (1998) guidelines cited by AFFA (2001) also include ethylene oxide. AFFA (2001)
notes that no studies could be found to demonstrate efficacy of ETO on its own, but that it was
effective in combination with methyl bromide (1:1.44 w/w) after 24 hours at 20-25oC.
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De Vos (1994) states that even products made after tanning and curing of hides cannot be
considered safe. AQIS (2001) has concluded, however, that the liming/dehairing process of
commercial tanning should destroy anthrax spores.
Semen/embryos: No references were found to the presence of anthrax in semen. It seems
unlikely that semen would be collected from a bull during the viraemic period.
Faeces: The faeces of infected animals may contain anthrax spores. De Vos (1994) advises
disposal of excreta by incineration or by burial in a 2 metre deep hole, covered with one part of
chloride of lime (at least 25% active chlorine) to three parts soil prior to filling in the hole.
Conclusions:
•
The body of an animal with anthrax contains vast numbers of anthrax bacteria.
The bacteria can survive for only a few days in an intact carcase. However, on
exposure to air, anthrax bacteria produce spores, which are very hardy and can
survive in the environment for many years.
•
Opening the carcase must be avoided at all costs.
•
Burning is the preferred method of carcase disposal. When handling infected
carcases it is important to avoid rupturing a carcase and/or spillage of body fluids.
•
Surface decomposition is a less desirable option. Carcases managed this way must
be sprayed with 5% formaldehyde and securely protected from scavengers.
•
Deep burial is the least desirable disposal option, because anthrax spores can
survive in the soil for decades. If carcases are deep buried, they must be at least two
metres beneath the surface and liberally covered with quicklime.
•
Skins and hides from anthrax infected animals are a significant disease risk. They
should be disposed of in a similar maner to fresh carcases.
•
Meat from animals clinically healthy at ante-mortem inspection is a negligible
transmission risk. No special disposal procedures are warranted.
•
Milk from affected herds should be pasteurised.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
Bengis, R.C. 1997, ‘Animal health risks associated with the transportation and utilisation of
wildlife products’, Rev sci tech Off int Epiz, vol. 16, pp. 104-110.
Bohm, R. 1990, ‘Resistance, survival, sterilisation and disinfection of spores of Bacillus
anthracis’, Salisbury Med Bull, vol. 68 (special supp), pp. 99-101, cited in de Vos 1994.
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Buxton, A., & Fraser, G. 1977, ‘Bacillus’, in Animal microbiology, eds. Buxton, A., & Fraser, G.,
Blackwell Scientific Publications, Oxford, cited in Bengis 1997.
de Vos, V. 1994, ‘Swine vesicular disease’, in Infectious diseases of livestock, Vol II, eds.
Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 12621289.
De Kock, G.vd W., Sterne, M., & Robinson, E.M. 1940, ‘Sterilisation of bone meal’, J Sth Afr vet
med Assoc, vol. 11, pp.138-141, cited in de Vos 1994.
Dragon, D.C., & Rennie, R.P. 1995, ‘The ecology of anthrax spores: tough but not invincible’,
Can Vet J, vol. 36, pp. 295-301.
Minett, F.C. 1950, ‘Sporulation and viability of B. anthracis in relation to environmental
temperature and humidity’, J Comp Pathol, vol. 60, pp. 161-176, cited in de Vos 1994.
Radostits, O.M., Gay, C.G., Blood, D.C., & Hinchcliff, K.W. (eds) 2000, Veterinary medicine,
ninth edition, W. B. Saunders Company Ltd, London.
Stein, C.D. 1947, ‘Some observations on the tenacity of Bacillus anthracis’, Vet Med, pp. 13-22,
cited in de Vos (1994) and Dragon and Rennie (1995).
Turnbull, P.C.B., Lindeque, P.M., Le Roux, J., Bennett, A.M., & Parks, S.R. 1998, ‘Airborne
movement of anthrax spores from carcass sites in the Etosha National Park, Namibia’, J Appl
Microbiol, vol. 84, pp. 667-676.
Whitford, H.W. 1978, ‘Factors affecting the laboratory diagnosis of anthrax’, J Am Vet Med
Assoc, vol. 173, pp.1467-1469, cited in de Vos 1994.
Whitford, H.W. 1987, A guide to the diagnosis, treatment and prevention of anthrax, World
Health Organization, cited in de Vos 1994.
WHO (World Health Organization) 1994, ‘Anthrax control and research, with special reference
to national programme development in Africa: memorandum from a WHO meeting’, Bulletin of
the World Health Organization, vol. 72, pp. 1-9.
WHO (World Health Organization) 1998, ‘Guidelines for the surveillance and control of anthrax
in humans and animals’, WHO/EMC/ZDI/98.6, cited in AFFA 2001.
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Heartwater
Agent:
Cowdria ruminantium
Agent type:
Bacterium
Persistence and inactivation:
Cowdria ruminantium is a rickettsia. It is an obligate intracellular parasite, infecting endothelial
cells of blood vessels of various organs, and is seen in the blood in association with red blood
cells, neutrophils and plasma. C. ruminantium is transmitted between vertebrate hosts by ticks of
the genus Amblyomma (Bezuidenhout et al 1994). Heartwater is not contagious, and even
experimental transmission by needle is difficult (Uilenberg pers comm).
The only literature of relevance pertains to work attempting to preserve the organism for
experimental purposes. Such work has generally shown that C. ruminantium is heat labile and
loses its viability within 12-38 hours at room temperature (Uilenberg pers comm). Alexander
(1931) was able to maintain C. ruminantium in defibrinated blood for up to 38 hours. He
observed that C. ruminantium generally did not survive for more than 24 hours at room
temperature, and in some cases lost viability within 12 hours.
In one exceptional case, infectivity was maintained in blood stored at room temperature for 4
days (Henning 1956).
Alexander (1931) noted that low temperatures seemed to favour the viability of C. ruminantium.
Infected ovine spleen and blood homogenates kept at -76oC maintained infective virus for at least
2 years (Neitz 1968). Logan (1987) reported effective preservation of C. ruminantium at -70oC to
-196oC for indefinite periods of time in a variety of organ suspensions. Blood collected from
infected goats and stored for as long as 72 hours at 4 degrees were still infectious to mice.
No reports were found on the survival of Cowdria ruminantium in carcases or animal products.
As Uilenberg (pers comm) has pointed out, this is partly due to the extreme difficulty in proving
whether or not the organism is alive.
Conclusions:
•
C. ruminantium is an obligate intracellular parasite that is spread by ticks.
•
Carcases and animal products pose a negligible transmission risk. No special
disposal precautions are warranted.
References:
Alexander, R. A., 1931, ‘Heartwater. The present state of our knowledge of the disease’, 17th
Report of the Director of Veterinary Services and Animal Industry, Union of South Africa, pp. 89150, cited by Logan (1987).
Bezuidenhout, J.D., Prozesky, L., du Plessis, J.L.,& van Amstel, S.R., 1994, ‘Heartwater’, in
Infectious diseases of livestock, Vol I, eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C.,
Oxford University Press, Cape Town, pp. 351-370.
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Henning, M.W., 1956,in Animal diseases in South Africa, third edition, Central News Agency
Limited, South Africa, pp. 1115-1179, cited in Logan (1987).
Logan, L. L., 1987, ‘Cowdria ruminantium: stability and preservation of the organism’,
Onderstepoort J Vet Res, vol. 54, pp. 187-191.
Neitz, W.O. 1968, ‘Heartwater’, Bull Off int Epiz, vol. 70, pp. 329-336, cited in Bezuidenhout et
al (1994).
Experts contacted:
Dr Gerrit Uilenberg
‘A Surgente’, route du Port
20130 Cargèse (Corsica)
FRANCE
P +33 4 9526 4083
[email protected]
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Brucellosis (B. abortus)
Agent:
Brucella abortus
Agent type:
Bacterium
Persistence and inactivation:
General characteristics: Huddleson (1943) reported that Brucella abortus may survive in the
environment for:
• up to eight months in aborted foetuses (in the shade)
• 2-3 months in wet soil
• 1-2 months in dry soil
• 3-4 months in faeces.
AFFA (2001) quotes a survival time for B. abortus of up to three weeks in the environment in
moist, humid conditions. Radostits et al (2000) describe infectivity as persisting for up to 100
days in winter and up to 30 days in summer in temperate climates. B. abortus has been found to
be quite resistant to a decrease in pH (Davies and Casey 1973), but sensitive to heat, sunlight, and
standard disinfectants, including phenolics, halogens, quaternary ammonium compounds, and
aldehydes at 0.5-1.0%. The bacterium can be maintained indefinitely by freezing (Radostits et al
2000).
B. abortus is excreted in greatest quantities in uterine discharges, abortions and foetal
membranes, and in milk. Urine, faeces and semen also contain the bacterium, as do hygromas
caused by the infection, but these sources are less important in the epidemiology of the disease.
Entry to the body is through ingestion, inhalation, via the conjunctiva, skin abrasions or intact
skin, and congenitally. Transmission is by direct contact or via the environment (Bishop et al
1994).
Carcases and meat products: B. abortus can be isolated from many organs of infected cattle,
creating a zoonotic risk for people handling carcases (Radostits et al 2000). Bishop et al (1994)
recommends incineration of foetuses, placenta and discharges from infected animals.
Apart from Huddleson’s (1943) report of persistence of B. abortus in aborted foetuses for up to
eight months in the shade, no references were found on the persistence or inactivation of in
carcases or meat products. AQIS (1999a) lists bovine brucellosis among those diseases “unable
to be transmitted via meat or meat products”.
Milk and milk products: Infected animals may shed B. abortus in colostrum and milk
intermittently throughout the lactation period. Unpasteurised milk has been responsible for
human infections (Bishop et al 1994).
Brucella spp are destroyed by pasteurisation (AQIS 1999b, Davies and Casey 1973, Keogh
1971). Heat treatment to ‘thermise’ milk for cheese production (62oC for 15 seconds) is not
sufficient to inactivate Brucella (AQIS 1999b). Davies and Casey (1973) inactivated B. abortus
in milk by heating for 15 seconds at 71.7oC, and at all time / temperature combinations down to 5
seconds / 65oC. Survival in whey was less than 4 days at 17-24oC (associated with a sharp
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decline in pH from 5.9 to 4.0), but at least 6 days at 5oC in whey (where pH dropped only to 5.4).
When B. abortus was stored in a citrate/phosphate buffer solution, viable organisms were found
after eight days at pH 4.0, while at pH <4.0 all bacteria died within 78 hours.
It is well documented that B. abortus survives the ordinary cheese making process and persists for
long periods in various cheese types. However, human infection from contaminated cheese is
rare (Keogh 1971).
Skins, hides and fibres: Contamination of these products may occur but in a draft assessment,
AFFA (2001) has determined that they do not pose a quarantine risk for B. abortus.
Semen/embryos: Brucella abortus can be found in the testicles, seminal vesicles and semen of
infected bulls (Radostits et al 2000). AQIS (1999c) states that B. abortus can survive freezing in
semen and that it can be transmitted by artificial insemination.
B. abortus is likely to be collected with uterine flushes containing embryos (AQIS 1999c). Six
washes will generally ensure the removal of the bacteria from the embryos (Stringfellow et al
1984), unless the zona pellucida is damaged (Stringfellow et al 1986). Embryo transfer probably
does not present a significant risk in the transmission of B. abortus (Campo et al 1987).
Faeces: B. abortus is excreted in the faeces (Bishop et al 1994), and additional contamination of
faeces is likely from vaginal discharge (Verger 1980).
It has been found that increasing the temperature of storage decreased the survival time of
Brucella spp in manure. Reported survival times include:
• 385 days at 8oC
• at least 8 months at 12oC
• 29 days at 25oC
• less than one day at 37oC
• less than 4 hours at around 70oC
(King 1957, Kuzdas and Morse 1954, Plommet 1977, Verger 1980).
Verger (1980) concluded that composting of bedding, faeces and urine would inactivate Brucella
within an hour. The organism would however survive long periods in a slurry and would be a
source of further infection unless treated with xylene 1000ppm.
Conclusions:
•
B. abortus is a sensitive bacterium that, under favourable conditions, can survive in
the environment for several weeks to months. It is readily inactivated by heat, UV
light and commonly used disinfectants.
•
B. abortus is excreted in all body fluids, with the greatest number of organisms in
uterine discharges, aborted foetuses, foetal membranes and milk. Infection occurs
with ingestion, inhalation or contact with the conjunctiva or abraded skin.
•
B. abortus was eradicated from Australia with a program that focused on the
diagnosis of infected cattle, with little emphasis on carcase disposal or animal
products.
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•
Carcases and animal products are not important in the epidemiology of the disease.
However, they are important from a human health perspective – most human cases
occur as a result of handling aborted foetuses or foetal membranes, or from
consuming unpasteurised milk or milk products from infected cattle.
•
Meat derived from affected herds is a negligible transmission risk.
•
Milk from affected herds should be pasteurised.
•
Semen should be destroyed. Embryos should be washed according to IETS
standards.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999a, Importation of sausage casings into
Australia, import risk analysis, December 1999, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999b, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999c, Import risk analysis report on the
importation of bovine semen and embryos from Argentina and Brazil into Australia, November
1999, AQIS, Canberra.
Bishop, G.C., Bosman, P.P., & Herr, S. 1994, ‘Bovine brucellosis’, in Infectious diseases of
livestock, Vol II, eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press,
Cape Town, pp. 1053-1066.
Campo, L.R., Tamayo, R., & Campo, C.H. 1987, ‘Embryo transfer from brucellosis-positive
donors: A field trial’, Theriogenology, vol. 27, p. 221, cited in Bishop et al 1994.
Davies, G., & Casey, A. 1973, ‘The survival of Brucella abortus in milk and milk products’, Br
vet J, vol. 129, pp. 345-353.
Huddleson, I.F. 1943, Brucellosis in man and animals, The Commonwealth Fund, New York,
cited in Bishop et al 1994.
Keogh, B.P. 1971, ‘Reviews of the progress of dairy science. Section B. The survival of
pathogens in cheese and milk powder’, J Dairy Res, vol. 38, pp. 91-111.
King, N.B. 1957, ‘The survival of Brucella abortus (USDA strain 2308) in manure’, J Am Vet
Med Ass, vol. 131, pp. 349-352, cited in Verger 1980.
Kuzdas, C.D., & Morse, E.V. 1954, ‘The survival of Brucella abortus, USDA strain 2308, under
controlled conditions in nature’, Cornell Vet, vol. 44, pp. 216-228, cited in Verger 1980.
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Plommet, M. 1977, ‘Studies on experimental brucellosis in cows in France’, in Bovine
brucellosis: an international symposium, College Station, London, eds. Crawford, R.P., &
Hidalgo, R.J., Texas A & M University Press, cited in Bishop et al 1994.
Radostits, O.M., Gay, C.G., Blood, D.C., & Hinchcliff, K.W. (eds) 2000, Veterinary medicine,
ninth edition, W. B. Saunders Company Ltd, London.
Stringfellow, D.A., Scanlon, C.M., Brown, R.R., Meadows, G.B., Gray, B.W., & Young White,
R. R. 1984, ‘Culture of bovine embryos after in vitro exposure to Brucella abortus’,
Theriogenology, vol. 21, pp. 1005-1012, cited in AQIS 1999c.
Stringfellow, D.A., Wolfe, D.F., Lauerman, L.H., & Sparling, P.H. 1986, ‘Resistance of
preimplantation bovine embryos to infection with Brucella abortus’, Am J Vet Res, vol. 47, pp.
1924-1927, cited in AQIS 1999c.
Verger, J. M. 1980, ‘Prevalence and survival of Brucella species in manures’, in Communicable
diseases resulting from storage, handling, transport and landspreading of manures. Proceedings
of a CEC workshop held at Hannover, November 1993, pp. 157-160.
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Brucellosis (B. melitensis)
Agent:
Brucella melitensis
Agent type:
Bacterium
Persistence and inactivation:
General characteristics: Survival of Brucella melitensis in the environment is similar to that of B.
abortus (Herr 1994). B. abortus has been reported to last up to several months in the
environment, longer in moist, cool conditions out of direct sunlight (see Bishop et al 1994,
Radostits et al 2000). Geering et al (1995) state that B. melitensis can survive for up to three
months in soil protected from sunlight and up to six months in necrotic placenta and foetus.
According to Mitscherlich and Marth (1984), B. melitensis is rapidly inactivated by moist heat
and outside pH range 5-8. The organism was inactivated at:
• between 7.5 and 10 minutes at 60oC
• between 5 and 7.5 minutes at 61.1oC
• less than 5 minutes at 62.8oC.
El-Daher et al (1990) found that in broth, B. melitensis survived for:
• more than 4 weeks at pH 5.5 or above
• less than three weeks at pH 5
• one day at pH 4
and did not survive below pH 4.
After an initial bacteraemia, B. melitensis localises in lymph nodes, udder and uterus or testes.
Infected females shed large quantities of bacteria in genital discharges and abortions. These
provide the main source of infection, which takes place principally via inhalation, but also via
abraded skin in goats and sheep. Shedding also occurs in milk, urine and semen. Ingestion of
infected milk or meat is a common means of transmission to humans (Garin-Bastuji et al 1998,
Geering et al 1995, Herr 1994).
Carcases and meat products: No specific references were found on the survival of B. melitensis in
carcases or meat products. B. abortus has been shown to survive up to 44 days in guinea pig
carcases in cold conditions (Timoney et al 1988). B. melitensis can survive for up to six months
in necrotic foetal and placental material (Geering et al 1995). Alton (1987) recommended that all
membranes and aborted foetuses should be incinerated where possible and if not buried deeply.
Humans have become infected with B. suis by ingestion of uncooked meat or bone marrow from
infected animals (Acha and Szyfres 1987). MacDiarmid and Thompson (1997) note that
infection could theoretically be picked up from infected carcases of sheep or goats by dogs or
pigs, but that pigs are dead-end hosts and transmission from dogs to other animals is rare.
Milk and milk products: B. melitensis is excreted in the milk of infected sheep and goats (Herr
1994) and milk products have been implicated by several authors as the source of human disease
(e.g. Thapar and Young 1986).
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Brucella spp are destroyed by pasteurisation (AQIS 1999, Davies and Casey 1973, Keogh 1971).
El-Daher et al (1990) found that survival of B. melitensis in dairy products was inversely
proportional to pH. They concluded that soft cheese was the most likely of the products
examined to present an infective risk, because it took up to 72 hours to fall below pH 4 and had
the highest bacterial counts at 48 hours. Yoghurt posed an intermediate risk, while bacteria were
not found in any of the milk samples by 24 hours. B. melitensis did not survive for even four
hours in buttermilk, which had an initial pH below 4.
In a review of the literature, Rammel (1967) noted that B. melitensis had been found in Feta after
4-16 days and up to 90 days in Pecorino cheeses.
Skins, hides and fibres: No specific references were found on this aspect of the disease in the
literature. Contamination of these products with genital discharges could be expected to occur.
In a draft assessment, AFFA (2001) has concluded that skins, hides and fibres could carry
Brucella for up to three months and that unprocessed product from endemic areas posed some
quarantine risk. Normal processing methods of alkaline scouring, acid pickling or liming /
dehairing should eliminate this risk in processed or partially-processed skins, hides or fibres.
Semen/embryos: B. melitensis is commonly shed in semen, although this seems only to have been
confirmed recently (Garin-Bastuji et al 1998). No reports were found of transmission of B.
melitensis from goat or sheep semen, nor of the infection of embryos by the agent. However,
AQIS (2000) has concluded that the quarantine risk posed by semen and embryos for B.
melitensis is high.
Faeces: Excretion of B. melitensis in faeces is not described in texts on the disease, although B.
abortus is shed by this route (Bishop et al 1994). However, faeces is highly likely to become
contaminated by the heavy load of organisms in vaginal discharges after parturition or abortion
(Verger 1980).
Survival characteristics of B. melitensis in faeces are likely to be similar to those of B. abortus.
Verger (1980) has shown that composting (to reach 70oC) of faeces, urine and straw bedding will
kill Brucella species within one hour. Studies showing long term survival of B. abortus in faecal
slurry in storage pits suggest that B. melitensis would show good viability in these conditions.
Xylene 1000ppm is an effective disinfectant for B. abortus (Verger 1980).
Conclusions:
•
B. melitensis is a relatively sensitive bacterium that, under favourable conditions,
can survive in the environment for several weeks to months. It is readily inactivated
by heat, UV light and commonly used disinfectants.
•
B. melitensis is excreted in all body fluids, with the greatest number of organisms in
uterine discharges, aborted foetuses, foetal membranes and milk. Infection occurs
with inhalation, ingestion or contact with the conjunctiva or abraded skin.
•
Carcases and animal products are not important in the epidemiology of the disease.
However, they are important from a human health perspective – most human cases
occur as a result of handling aborted foetuses or foetal membranes, or from
consuming unpasteurised milk or milk products from infected sheep or goats.
•
Meat derived from affected herds is a negligible transmission risk.
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•
Milk from affected herds should be pasteurised.
•
Semen and embryos should be destroyed.
References:
Acha, P.N., & Szyfres, B. 1987, Zoonoses and communicable diseases common to man and
animals, second edition, Pan American Health Organisation, Washington, 963 pp., cited in
MacDiarmid and Thompson 1997.
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
Alton, G.G. 1987, ‘Control of Brucella melitensis infection in sheep and goats – a review’, Trop
Anim Hlth Prod, vol. 19, pp. 65-74.
AQIS (Australian Quarantine and Inspection Service) 1999, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 2000, An analysis of the disease risks, other
than scrapie, associated with the importation of ovine and caprine semen and embryos from
Canada, the United States of America and member states of the European Union, final report,
AQIS, Canberra.
Bishop, G.C., Bosman, P.P., & Herr, S. 1994, ‘Bovine brucellosis’, in Infectious diseases of
livestock, Vol II, eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press,
Cape Town, pp. 1053-1066.
Davies, G., & Casey, A. 1973, ‘The survival of Brucella abortus in milk and milk products’, Br
vet J, vol. 129, pp. 345-353.
El-Daher, N., Na’was, T., & Al-Qaderi, S. 1991, ‘The effect of the pH of various dairy products
on the survival and growth of Brucella melitensis’, Annals Trop Med Parasitol, vol. 84, pp. 523528.
Garin-Bastuji, B., Blasco, J. M., Grayon, M., & Verger, J. M., 1998, ‘Brucella melitensis
infection in sheep: present and future’, Vet. Res., vol. 29, pp. 255-274 cited by AQIS (2000).
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
Herr, S. 1994, ‘Brucella melitensis infection’, in Infectious diseases of livestock, Vol II, eds.
Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 10731075.
Keogh, B.P. 1971, ‘Reviews of the progress of dairy science. Section B. The survival of
pathogens in cheese and milk powder’, J Dairy Res, vol. 38, pp. 91-111.
MacDiarmid, S.C., & Thompson, E.J. 1997, ‘The potential risks to animal health from imported
sheep and goat meat’, Rev sci tech Off int Epiz, vol. 16(1), pp.45-56.
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Mitscherlich, E., & Marth, E.H. 1984, in Microbial survival in the environment, Springer-Verlag,
Berlin, cited in AFFA 2001.
Radostits, O.M., Gay, C.G., Blood, D.C., & Hinchcliff, K.W. (eds) 2000, Veterinary medicine,
ninth edition, W. B. Saunders Company Ltd, London.
Rammell, C.G. 1967, Aust J Dairy Technol, vol. 22, p. 40, cited in Keogh 1971.
Thapar, M.K., & Young, E.J. 1986, ‘Urban outbreak of goat cheese brucellosis’, Pediatric Inf
Dis, vol. 5, pp. 640-643, cited by El-Daher et al 1991.
Timoney, J.F., Gillespie, J.H., Scott, F.W., & Barlough, J.E. (eds) 1988, Hagan and Bruner’s
microbiology and infectious diseases of animals, Comstock Publishing Associates, Ithaca, 951
pp., cited in MacDiarmid and Thompson 1997.
Verger, J. M. 1980, ‘Prevalence and survival of Brucella species in manures’, in Communicable
diseases resulting from storage, handling, transport and landspreading of manures. Proceedings
of a CEC workshop held at Hannover, November 1993, pp. 157-160.
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Bovine tuberculosis
Agent:
Mycobacterium bovis
Agent type:
Bacterium
Persistence and inactivation:
General characteristics: Mycobacterium bovis is relatively resistant to heat, desiccation, and
disinfectants (Radostits et al 2000). Heat treatments (hot air, burning, cooking, pasteurisation,
pressurised steam) are the most useful methods for inactivating M. bovis.
M. bovis is relatively resistant to chemical disinfectants by virtue of its waxy, hydrophobic cell
wall (Russell 1996). Inorganic acids, alkalis, quaternary ammonium compounds and chlorides
are ineffective. Formalin (3%), lysol (2%), phenol (2.5%), activated chloramine (1-3%), cresols
and iodophors are effective (Huchzermeyer et al 1994).
A number of studies have been conducted on the survival of M. bovis in the environment, and the
findings are reasonably consistent.
An Australian study demonstrated survival of M. bovis for less than eight weeks in the shade and
less than four weeks in direct sunlight (Duffield and Young 1985). Tanner and Michel (1999), in
a study in the Kruger National Park in South Africa, found that M. bovis survived for up to four
weeks in spiked faeces and up to six weeks in buffalo lung or lymph node. Moisture and
temperature were the major determinants of viability. Ultraviolet light was not thought to be a
factor unless there was a lack of moisture, with maximum survival in faeces in sunny but moist
conditions.
In New Zealand, Jackson et al (1995) looked at the survival of M. bovis adsorbed onto cotton
strips and placed in different natural habitats. Maximum survival was found in possum dens (1428 days in winter and spring, less than seven days in summer), then on the forest floor (14-28
days in winter and spring, less than four days in summer). No bacteria were found on pasture
after four days.
The authors concluded that M. bovis does not survive long outside hosts, and that environmental
contamination may therefore be relatively unimportant in the epidemiology of the disease
(Jackson et al 1995).
M. bovis may be localised in the body, or it can be widely disseminated. Primary foci are usually
in the respiratory and/or gastrointestinal tracts (Huchzermeyer et al 1994). Transmission of
bovine tuberculosis is almost exclusively via the respiratory route, although infection can occur
via alimentary, congenital, cutaneous, and venereal routes, as well as through the teat canal.
Mycobacterium bovis is present in respiratory aerosols and may also be shed in vaginal
secretions, milk, urine and faeces from cattle with generalised tuberculosis. Tuberculosis is
usually spread to humans through unpasteurised milk and dairy products (Huchzermeyer et al
1994, Scanlon and Quinn 2000b).
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Carcases and meat products:
Feeding of tuberculosis-infected carcases poses some infective risk, particularly through infected
lymph nodes (Huchzermeyer et al 1994). Meat harbours few or no tubercle bacilli, and the oral
infective dose is large in comparison with the respiratory infective dose, leading Francis (1973) to
conclude that the risk posed to humans by eating tuberculous meat was slight.
In Tanner and Michel’s (1999) study in the Kruger National Park, M. bovis from naturally
infected buffalo survived for up to six weeks in lung and lymph nodes in the winter and 5-14 days
in the other seasons. The authors noted that previous investigations of M. bovis in possum and
badger carcases produced similar results, showing survival times of 2-4 weeks. Inactivation is
aided by the decomposition of the carcase, and by the scavenging of vertebrates, insects and
helminths.
Maximum survival for M. bovis buried in moist soil was five days. This was compared to the
findings of O’Reilly and Daborn (1995), who were unable to isolate M. bovis from three buried
badger carcases after two, three, and six weeks respectively.
Merkal & Whipple (1980) found that inactivation of M. bovis in meat products by heat required
time / temperature combinations 6-7oC below those formerly identified for the M. avium – M.
intracellulare complex. A one log reduction required approximately 25 minutes / 53oC to one
minute / 61oC. A five log reduction required 350 minutes / 53oC to one minute / 68oC (all
approximate figures read from graph). Ultraviolet radiation, amphyl and formaldehyde vapour
were found separately and in any combination to destroy M. bovis in thin meat emulsion smears.
Alkaline hydrolysis was found to completely inactivate M. bovis BCG in a study by Kaye et al
(1998). This experiment involved the placement of 114-136 kg loads of animal carcases into an
animal digester, with pure cultures of various micro-organisms in dialysis bags also placed in the
digester. The carcases were covered by hot alkaline solution and kept at 110-120oC for 18 hours
in the digester. The method was proposed as an alternative to incineration.
Milk and milk products: Large numbers of infective doses of M. bovis may be shed in the milk of
infected cows (Huchzermeyer et al 1994). Drinking infected milk is a common method of spread
in young animals (Radostits et al 2000).
Grant et al (1996) demonstrated inactivation of M. bovis in milk by heating at 63.5oC for 20
minutes, ten minutes within the specifications of holder or low-temperature, long time (LTLT)
pasteurisation. Initial inocula were at a high level (107 cfu/mL). M. bovis exhibited a linear
thermal death curve and was second in heat sensitivity only to M. fortuitum of the five
mycobacteria tested.
Harrington and Karlson (1965) have also shown that M. bovis does not survive LTLT or hightemperature, short time (HTST, 71.1oC /15 seconds) pasteurisation in skim milk. Huchzermeyer
et al (1994), however, state that HTST pasteurisation is not always effective in destroying the
bacilli, even when the time is extended to 20-25 seconds. The note of warning is a reminder that
milk may contain a high number of cells or pus, which will reduce the efficacy of the
pasteurisation in a way not observed by the in vitro studies.
If milk is inadequately pasteurised and made into sour milk, buttermilk, yoghurt and cream
cheese, these milk products may contain M. bovis for up to 14 days after their preparation. In
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butter the bacterium may survive for 100 days (Huchzermeyer et al 1994). The pH of sour milk
does not destroy tubercle bacilli (Mattick and Hirsch 1946).
AQIS (1999a) gives examples of survival times for a range of cheeses:
• hard cheese 5-30 days
• semi-soft cheese 305 days
• camembert style soft cheese 47 days.
Keogh (1971) also reviewed survival times of M. bovis in a range of cheeses, and concluded that
the requirement for minimum maturing periods in some countries (e.g. 60, 90 or 120 days before
sale) could not be justified, because the survival time is so highly variable and can often exceed
these periods. There was however a strong case for pasteurisation of all milk for cheese
manufacturing.
Skins, hides and fibres: No specific reports were found on this aspect of the disease. M. bovis
might be expected to found as a superficial contaminant of skins, hides and fibres, but its survival
in the absence of moisture would be short. In a draft assessment, AFFA (2001) has concluded
that these products do not pose a quarantine risk for bovine tuberculosis.
Semen/embryos: M. bovis may be present in the semen of infected bulls, originating from
tuberculous lesions in the prepuce or when phagocytes containing M. bovis facilitate movement
of the organism into the semen (Thoen et al 1977). M. bovis can survive in frozen semen.
Venereal infection may occur by semen infected by both routes. The risk of transmission from
infected bulls has been assessed as moderate (AQIS 1999b).
There is a reasonable likelihood that uterine flushes containing embryos from infected donor
cows could contain M. bovis, as genital tuberculosis may occur in cows (AQIS 1999b). Rohde et
al (1990) found that washing of embryos did not always remove other Mycobacterium spp. Some
risk of transmission of M. bovis is therefore presumed despite washing of embryos.
Faeces: Scanlon and Quinn (2000a) describe numerous reports of M. bovis shed in the faeces of
tuberculous and reactor cattle. There is an infective risk to grazing cattle if the slurry is spread on
pasture (Jones 1980), although land used for tillage hay or silage is ideal for slurry from reactor
herds (Scanlon and Quinn 2000b). Mechanical agitation of slurry can also create aerosols that are
infective to animals or man (Scanlon and Quinn 2000a).
M. bovis that may survive for long periods in stored slurry and in the environment if spread on the
land (Russell 1996). Scanlon and Quinn (2000b) found that M. bovis survived for six months in
sterilised slurry in screw capped bottles, stored in the dark at an ambient temperature. Other
reports include Williams and Hoy (1930) who reported survival for 4 months in slurry stored in a
jar in an underground cellar, and Dokoupil (1964) who documented survival for 176 days in
slurry at 5oC.
Scanlon and Quinn (2000a) describe two techniques of reducing the bacterial numbers to
acceptable level. The first is long term storage, which is the slower but less expensive technique.
It may be necessary to store the slurry for up to 6 months before all the M. bovis bacteria are
inactivated naturally. The alternative is treat slurry with chemicals prior to spreading. Any
chemical used must retain its activity in the presence of large amounts of organic matter.
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The efficacy of five disinfectants against M. bovis were tested at varying concentrations in cattle
slurry in screw-tapped bottles. Acetone (22.5%) was effective within 24 hours, ammonium
hydroxide (1%) within 36 hours, and chloroform (0.5%), ethyl alcohol (17.5%), and xylene (3%)
within 48 hours (Scanlon and Quinn 2000a).
Conclusions:
•
Mycobacterium bovis is a sensitive organism which, under favourable conditions, can
survive in the environment for a few weeks.
•
Transmission between cattle is mainly by respiratory aerosol. Most human cases
occur as a result of consuming unpasteurised milk from infected cattle.
•
Fresh carcases should be disposed of in a manner that prevents scavenging.
•
Meat and skins from affected herds pose a negligible transmission risk. No special
disposal precautions are warranted.
•
Milk from affected herds should be pasteurised.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999a, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999b, Import risk analysis report on the
importation of bovine semen and embryos from Argentina and Brazil into Australia, November
1999, AQIS, Canberra.
Dokoupil, S. 1964, ‘Survival of M. tuberculosis in grass, soil, bedding in cow sheds and urine’,
Vedecke Prace Vyzkumneho Ustavu Veterinarniho Lekarstvi v Brne, vol. 3, pp. 49-52, cited in
Scanlon and Quinn 2000a.
Duffield, B. J., & Young, D. A. 1985, ‘Survival of Mycobacterium bovis in defined
environmental conditions’, Vet Microbiol, vol. 10, pp. 193-197, cited in Scanlon and Quinn
(2000b).
Francis, J. 1973, ‘Very small public health risk from flesh of tuberculous cattle’, Aust vet J, vol.
49, pp. 496-497, cited in MacDiarmid and Thompson 1997.
Grant, I.R., Ball, H.J., & Rowe, M. T. 1996, ‘Thermal inactivation of several Mycobacterium
spp. in milk by pasteurisation’, Letters in Appl Microbiol, vol.22, pp. 253-256.
Harrington, R. & Karlson, A. G. 1965, ‘Destruction of various kinds of mycobacteria in milk by
pasteurization’, Appl Microbiol, vol. 13, pp. 494-495, cited in Grant et al 1996.
Huchzermeyer, H.F.K.A., Bruckner, G.K., Van Heerden, A., Kleeberg, H.H., van Rensburg,
I.B.J., Koen, P., & Loveday, R.K. 1994, ‘Tuberculosis’, in Infectious diseases of livestock, Vol II,
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eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp.
1425-1444.
Jackson, R., de Lisle, G.W., & Morris, R.S. 1995, ‘A study of the environmental survival of
Mycobacterium bovis on a farm in New Zealand’, New Zealand Vet J, vol. 43, pp. 346-352.
Kaye, G.I., Weber, P.B., Evans, A., & Venezia, R.A. 1998, ‘Efficacy of alkaline hydrolysis as an
alternative method for treatment and disposal of infectious animal waste’, Contemp Topics Lab
Animal Sci, vol. 37(3), pp. 43-46.
Keogh, B.P. 1971, ‘Reviews of the progress of dairy science. Section B. The survival of
pathogens in cheese and milk powder’, J Dairy Res, vol. 38, pp. 91-111.
MacDiarmid, S.C., & Thompson, E.J. 1997, ‘The potential risks to animal health from imported
sheep and goat meat’, Rev sci tech Off int Epiz, vol. 16(1), pp.45-56.
Mattick, A.T.R., & Hirsch, A. 1946, ‘Sour milk and the tubercule bacillus’, Lancet, vol. 250, pp.
417-417, cited in AQIS 1999.
Merkal, R.S., & Whipple, D.L. 1980, ‘Inactivation of Mycobacterium bovis in meat products’,
Appl Env Microbiol, vol. 40, pp. 282-284.
O’Reilly, L.M., & Daborn, C.J. 1995, ‘The epidemiology of Mycobacterium bovis infections in
animals and man: a review’, Tubercle and Lung Disease, vol. 76 supplement 1, pp. 1-46, cited in
Tanner and Michel 1999.
Rohde, R.F., Shulaw, W.P., Hueston, W.D., Bech-Nielsen, S., Haibel, G.K., & Hoffsis, G.F.
1991, ‘Isolation of Mycobacterium paratuberculosis from washed bovine ova after in vitro
exposure’, Am J Vet Res, vol. 51, pp. 708-710, cited in AQIS 1999b.
Radostits, O.M., Gay, C.G., Blood, D.C., & Hinchcliff, K.W. (eds) 2000, Veterinary medicine,
ninth edition, W. B. Saunders Company Ltd, London.
Russell, A. D. 1996, ‘Activity of biocides against mycobacteria’, J Appl Bacteriol, vol. 81, pp.
87S-101S, cited in Scanlon and Quinn (2000a).
Scanlon, M.P., & Quinn, P.J. 2000a, ‘Inactivation of Mycobacterium bovis in cattle slurry by five
volatile chemicals’, J Appl Microbiol, vol. 89, pp.854-861.
Scanlon, M.P., & Quinn, P.J. 2000b, ‘The survival of Mycobacterium bovis in sterilized cattle
slurry and its relevance to the persistence of this pathogen in the environment’, Irish Vet J, vol.
53, pp. 412-415.
Tanner, M., & Michel, A.L. 1999, ‘Investigation of the viability of M. bovis under different
environmental conditions in the Kruger National Park’, Onderstepoort J Vet Res, vol. 66, pp.185190.
Thoen, C.O., Himes, E.M., Stumpff, C.D., Parks, T.W., & Sturkie, H.N. 1977, ‘Isolation of
Mycobacterium bovis from the prepuce of a herd bull’, Am J Vet Res, vol. 38, pp. 877-878, cited
in AQIS 1999b.
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Williams, R.S., & Hoy, W.A. 1930, ‘Survival and spread of pathogenic bacteria of veterinary
importance within the environment’, Vet Bull, vol. 45, pp. 543-550, cited in Scanlon and Quinn
(2000b).
Experts contacted:
Dr Debby Cousins
Principal Microbiologist
Australian Reference Laboratory for Bovine Tuberculosis
Agriculture WA
3 Baron-Hay Court
SOUTH PERTH WA 6151
P 08 9368 3429
F 08 9474 1881
[email protected]
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Haemorrhagic septicaemia
Agent:
Pasteurella multocida
Agent type:
Bacterium
Persistence and inactivation:
General characteristics: Haemorrhagic septicaemia has been thoroughly reviewed by De Alwis
(1999). Information on the survival of Pasteurellae under different conditions is scant (De Alwis,
pers comm), probably because the existence of a carrier state in animals makes eradication by
stamping out appear unfeasible. (Eradication has been attempted on Lombok, Indonesia, using
mass vaccination. The disease does not appear to have been eradicated (De Alwis 1999).)
De Alwis (1999) states that “In general, P. multocida does not survive long enough outside the
animal to become a significant source of infection, although survival may be longer in moist
conditions”. The organism has been reported to survive for 2-3 weeks in sterilised soil (Bain et al
1982), although it could not be recovered from artificially infected, sterilised earth and mud from
Malaysian rice fields after several hours exposure to sunlight (FAO 1959). Nor could it be
recovered after 24 hours from mud in which buffaloes wallow (Bain et al 1982).
Carcases and meat products: No specific references to this aspect of the disease were found in the
literature. It is believed that P. multocida can survive in animal tissues, including rotting
carcases, for a few days. Bacteria have been found at 10ll – 1012 CFU/mL of blood of carcases
20-21 hours after death. Carcases may be a source of infection during these few days, and the
dumping of carcases in rivers has been implicated in downstream spread of the disease (De Alwis
1999).
Milk and milk products: AQIS (1999a) have listed P. multocida (Asian strain, serotype B and
African form serotype E) as an “organism not considered to be a quarantine hazard in dairy
products”, noting that the agent is unlikely to be in the milk of animals producing milk for human
consumption. No references were found during this review to the presence of P. multocida in
milk.
Skins, hides and fibres: Skin is unlikely to be a source of infectivity, given that the organism
spreads via ingestion of feed and water contaminated with nasal discharge and saliva, and does
not survive more than few days in the environment. In an interim assessment, AFFA (2001) have
classified P. multocida as not presenting a quarantine hazard in skins and hides.
Semen/embryos: There are no reports of P. multocida having been recovered from bovine semen,
nor of haemorrhagic septicaemia being transmitted by artificial insemination in cows. However,
other serotypes of P. multocida have been isolated from the prepuce and urine of healthy dogs,
and these can be transferred from the dog to the bitch at mating. AQIS (1999b) have therefore
concluded that “it is likely that infected semen can transmit the infective organism to susceptible
cows via artificial insemination” and permit importation of bovine semen only from HS-free
countries. Similarly, there is a presumptive risk of P. multocida on embryos or in uterine flushes,
and the same quarantine restriction is imposed.
Faeces: No references were found in the literature to the excretion of P. multocida in faeces.
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Conclusions:
•
Pasteurella multocida is a sensitive bacterium which, under favourable conditions,
can survive in the environment for only a few days.
•
Transmission between cattle is mainly by respiratory aerosol.
•
Carcases and animal products pose a negligible transmission risk. No special
disposal precautions are warranted.
•
Semen should be destroyed.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999, Import risk analysis report on the
importation of bovine semen and embryos from Argentina and Brazil into Australia, November
1999, AQIS, Canberra.
Bain, R.V.S., De Alwis, M.C.L., Carter, G.R., & Gupta, B.K. 1982, Haemorrhagic septicaemia,
FAO Animal Production and Health Paper No 33, FAO, Rome, cited in De Alwis 1999.
Bredy, J.P., & Botzler, R.G. 1989, ‘The effects of six environmental variables on Pasteurella
multocida populations in water’, J Wildl Dis, vol. 25(2), pp. 232-239.
De Alwis, M.C.L. 1999, Haemorrhagic septicaemia, ACIAR Monograph No 57, Australian
Centre for International Agricultural Research, Canberra, 141 pp.
FAO (Food and Agriculture Organisation) 1959, Report of the FAO meeting on haemorrhagic
septicaemia, Manila, Philippines, Nov-Dec, FAO, Rome, cited in De Alwis 1999.
Experts contacted:
Dr Malcolm C. L. De Alwis
Veterinary Research Institute
PO Box 28
Peradeniya
SRI LANKA
[email protected],ac.lk
Professor Sam Maheswaran
Veterinary Pathobiology
205 Vet S
1971 Commonwealth Avenue
St Paul, Minnesota 55108
USA
P +1 612 625 6264
[email protected]
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Glanders
Agent:
Burkholderia mallei
Agent type:
Bacterium
Persistence and inactivation:
General characteristics: B. mallei may found in the urine, saliva, tears, faeces, nasal discharges
and pus of infected animals (Bishop 1994). Transmission is mostly via ingestion from
contaminated feed, water and utensils, and the cutaneous form of the disease (farcy) may arise
from contamination of wounds. Transmission via inhalation has been demonstrated but its
importance in the field is unknown (Radostits et al 2000).
B. mallei is relatively susceptible to environmental influences. Bishop (1994), citing other texts,
notes that the bacterium is destroyed by sunlight in 24 hours, and by most common disinfectants,
including phenol, formalin, chlorine, potassium permanganate and copper sulphate. B. mallei
retains infectivity for three to five weeks in damp media, for 20-30 days in decomposing material,
for about 20 days in clean water and for about six weeks in contaminated stables.
Carcases and meat products: No specific references to this aspect of the disease were uncovered
during this review. The appearance of glanders in lions in an Italian zoo has been attributed to the
feeding of contaminated meat from imported horses (Battelli et al 1973). Bishop (1994) notes
that, in South Africa, clinical cases of glanders must be destroyed and the carcases burnt or
buried.
Skins, hides and fibres: AFFA (2001) has determined that there is some risk of at B. mallei being
found on the unprocessed skins of equids. As the agent is a non-spore forming bacillus, it is
unlikely to survive the liming or acid pickling processes applied to skins.
Semen/embryos: No references were found on the presence of B. mallei in semen or on embryos.
Faeces: B. mallei may be found in the faeces of infected animals (Bishop 1994).
Conclusions:
•
B. mallei is a sensitive bacterium which, under favourable conditions, can survive in
the environment for a few weeks. It is readily destroyed by exposure to sunlight and
commonly used disinfectants.
•
B. malleis is excreted in nasal discharges, urine, saliva, faeces and pus from infected
animals. Infection occurs by ingestion or contamination of abraded skin.
•
Fresh carcases should be disposed of in a manner that precludes scavenging.
•
Meat and processed skins pose a negligible transmission risk. No special disposal
precautions are warranted.
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References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
Battelli, C., Contento, F., Corsalini, T., Goffredo, G., Lazari, P., Puccini, V., & Sobrero, L. 1973,
‘Glanders in a group of lions in captivity’, Veterinaria Italiana, vol. 24, pp. 87-112, 113-116,
cited in Bishop 1994.
Bishop, G.C. 1994, ‘Glanders’, in Infectious diseases of livestock, Vol II, eds. Coetzer, J.A.W.,
Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 1046-1050.
Radostits, O.M., Gay, C.G., Blood, D.C., & Hinchcliff, K.W. (eds) 2000, Veterinary medicine,
ninth edition, W. B. Saunders Company Ltd, London.
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Contagious equine metritis
Agent:
Taylorella equigenitalis
Agent type:
Bacterium
Persistence and inactivation:
General characteristics: Most, if not all, of the published information on the survival of T.
equigenitalis relates to its survival for diagnostic purposes. Sahu et al (1979) compared the
survival of the organism in exudate in the presence of three transport media, and in the absence of
a transport medium. In day one, bacterial numbers decreased 15-fold at 22oC and two-fold at
4oC, but did not deteriorate at -70oC, in the absence of medium. Amies with charcoal was
generally better than modified Amies without charcoal or Stuart’s, with 19% of Taylorella
surviving to day 10 at the three temperatures. Studies by Timoney et al (1979) showed the
organism to be relatively heat labile, with thermal death times for pure culture in vaginal
discharge ranging from at least 27 minutes at 40oC to less than one minute at 50oC. Sahu and
Dardiri (1980) showed T. equigenitalis to be particularly susceptible to pH below 4.5.
Heath (pers comm) advised that “outside the body the organism would not be long-lived and in
exudate alone at ambient temperature would most likely be dead within 10-14 days, maybe
shorter – my opinion only. Chilling increases survival time but the role of a good transport
medium is important to survival over time”. T. equigenitalis in dried exudates is susceptible to
ten minutes exposure to chlorhexidine diacetate (2%) or alkyldimethylbenzylammonium chloride
(10%) (Henton 1994).
Carcases and meat products: No studies on the survival of T. equigenitalis in carcases were
uncovered during this review. (Three independent experts reported a similar lack of information
in the literature.) Heath (pers comm) speculated that faster growing commensal aerobes and
anaerobes, as well as low pH, would inhibit the growth of Taylorella. Infection is limited to the
reproductive tract of mares and surface contamination of the external genitalia of stallions
(Timoney 1996).
Skins, hides and fibres: The risk of CEM infectivity via skins or hides should be negligible, given
the normal route of transmission.
Semen/embryos: CEM can be transmitted by artificial insemination (Timoney 1996). The
organism has frequently been isolated from the terminal urethra of carrier stallions, and less
commonly from the pre-ejaculatory fluid of such animals. There is also the risk of indirect
contamination of semen via materials and equipment carrying the organism. Timoney (pers
comm) believes that provided high standards of breeding shed management are upheld, semen
from non-infected stallions from CEM-infected premises can be used safely. However,
decontamination of semen from carrier stallions using antimicrobials has not been successfully
demonstrated (Timoney et al 1979).
Faeces: Any T. equigenitalis found in faeces would be an incidental contaminant. It is unlikely
the organism would be found in significant numbers in faeces, or that it would pose any threat of
infectivity, given the route of transmission.
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Conclusions:
•
T. equigenitalis is a fragile bacterium which, under normal conditions, can survive
no longer than a week or two in the environment.
•
Disease spread occurs by venereal means.
•
Carcases and animal products pose a negligible transmission risk. No special
disposal precautions are warranted.
•
The semen from infected stallions should be destroyed.
References:
Henton, M.M. 1994, ‘Taylorella equigenitalis infection’, in Infectious diseases of livestock, Vol II,
eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp.
1510-1513.
Sahu, S.P., Dardiri, A.H., Rommel, F.A., & Pierson, R.E. 1979, ‘Survival of contagious equine
metritis bacteria in transport media’, Am J Vet Res, vol. 40, pp. 1040-1042.
Sahu, S.P., & Dardiri, A.H. 1980, ‘Contagious equine metritis: isolation and characterization of
the etiologic agent’, Am J Vet Res, vol. 41, pp. 1379-1382.
Timoney, P.J. 1996, ‘Contagious equine metritis’, Comp Immun Microbiol infect Dis, vol. 19(3),
pp. 199-204.
Timoney, P.J., O’Reilly, P.J., Harrington, A.M., McCormack, R., & McArdle, J.F. 1979,
‘Survival of Haemophilus equigenitalis in different antibiotic-containing semen extenders’, J
Reprod Fert, Suppl vol. 27, pp. 377-381.
Timoney, P.J., Ward, J., McArdle, J.F., & Harrington, A.M. 1979, ‘Thermal death times of the
organism of contagious equine metritis 1977’, Vet Rec, vol. 104, p. 530.
Experts contacted:
Dr Peter Heath
Veterinary Laboratories Agency
Rougham Hill, Bury St Edmunds
Suffolk IP33 2RX
UK
T +44 1284 724499
F +44 1284 724500
[email protected]
Dr Peter Timoney
Department of Veterinary Science
108 Gluck Equine Research Center
Lexington, Kentucky 40546-0099
USA
P +1 859 257 1531
F +1 859 257 8542
[email protected]
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Potomac fever
Agent:
Ehrlichia risticii
Agent type:
Bacterium
Persistence and inactivation:
Ehrlichia risticii is a rickettsia. It is an obligate intracellular parasite, seen principally in the
bloodstream in association with monocytes. The agent is also found in various tissues and organs
including small and large intestine and associated lymph nodes, and it is shed in faeces (Holland
1990, Radostits et al 2000).
The transmission of E. risticii has not been elucidated. It appears that the organism is not
contagious. According to Palmer (1987), “it is difficult if not impossible to transmit using the
faecal-oral route”, although Radostits et al (2000) cite reports of experimental transmission via
the oral route, acknowledging that the significance of this in the natural disease is unknown.
Infection can be artificially induced by injection of whole blood from an infected to a susceptible
horse (Jenny 1984). The natural means of transmission is thought to take place via a vector,
probably a tick. Transplacental infection has also been recorded (Holland 1990).
No reports were found on the survival of E. risticii in vitro, in carcases or in animal products.
Conclusions:
•
Ehrlichia risticii is an obligate intracellular parasite that is spread by ticks.
•
Carcases and animal products pose a negligible transmission risk. No special
disposal precautions are warranted.
References:
Holland, C.J. 1990, ‘Biologic and pathogenic properties of Ehrlichia risticii: the etiologic agent
of equine monocytic ehrlichiosis’, in Current topics in veterinary medicine and animal science.
Vol. 54. Ehrlichiosis: a vector-borne disease of animals and humans, eds. Williams, J.C., &
Kakoma, I., Kluwer Academic Publishers, Netherlands, pp. 68-77.
Jenny, A.L. 1984, ‘Potomac horse fever – National Veterinary Services Laboratories (USDA)
report’, Am Assoc Eq Practnr Newsltr, no. 2, pp. 64-66, cited in Holland 1990.
Palmer, J.E. 1987, ‘Potomac horse fever’, in Current therapy in equine medicine, ed. Robinson,
N.E., W.B. Saunders Company, Philadelphia, pp. 92-93.
Radostits, O.M., Gay, C.G., Blood, D.C., & Hinchcliff, K.W. (eds) 2000, Veterinary medicine,
ninth edition, W. B. Saunders Company Ltd, London.
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Contagious bovine pleuropneumonia
Agent:
Mycoplasma mycoides subsp mycoides SC (bovine type)
Agent type:
Mycoplasma
Persistence and inactivation:
General characteristics: M. mycoides subsp mycoides SC is reported to survive off the host for up
to three days in tropical climates and for up to two weeks in temperate zones (Schneider et al
1994). Laak (1992) states that pathology specimens may be stored at room temperature for weeks
or months without affecting recovery of the agent. It survived for 216 hours in the shade, and 168
hours in the open on inoculated hay, and was inactivated as a result of both desiccation and UV
light in a study by Windsor and Masiga (1977).
The organism is inactivated within 60 minutes at 50oC and within two minutes at 60oC, but may
persist for at least twelve months in frozen lung tissue (Schneider et al 1994). It is killed below
ph 5.5 (Windsor unpublished, cited in Windsor and Masiga 1977). The critical temperature for
survival of the V5 vaccine strain that was used in Australia is 45oC (Hudson 1968).
M. mycoides subsp mycoides SC is sensitive to 1% phenol (three minutes to inactivation), 0.05%
formaldehyde (30 seconds), and 0.01% mercuric chloride (one minute) (Provost et al 1987).
CBPP is spread by the inhalation of infected aerosols from the respiratory tract (Laak 1992) and
there is a possibility of spread via infected droplets of urine (Schneider et al 1994).
Carcases and meat products: M. mycoides has been shown to survive in placenta for 72 hours,
when it was overgrown by bacteria (Windsor and Masiga 1977). No specific references were
found during this review regarding the survival of Mycoplasma mycoides in carcases or meat.
Transmission by ingestion of ground-up infected lung has been demonstrated experimentally
(Hyslop 1959), although this study seems to have disappeared from most subsequent reviews.
Schneider et al (1994) state that “neither ingestion of infected fodder nor direct exposure of
diseased organs of animals suffering from CBPP, will cause transmission”. AQIS (1999b) has
determined that CBPP is unable to be transmitted by meat or meat products.
Milk and milk products: No references were found to the excretion of M. mycoides in milk.
AQIS (1999a) has concluded that there is little risk of introducing the agent in milk. Even if the
agent were present, experience with other Mycoplasma spp indicate that it should be inactivated
by pasteurisation.
Skins, hides and fibres: Skins, hides and fibres are unlikely to harbour infective agent for long
periods. In a draft assessment, AFFA (2001) has concluded that these products do not pose a
quarantine risk for CBPP.
Semen/embryos: No reports were uncovered during this review of Mycoplasma mycoides
appearing in semen or embryos.
Faeces: Mycoplasma mycoides could be expected to be a contaminant of faeces. The agent is
found in the urine of infected animals (Windsor and Masiga 1977).
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Conclusions:
•
M. mycoides is a sensitive organism which, under favourable conditions, may survive
in the environment for up to two weeks.
•
Transmission between cattle is by respiratory aerosol.
•
Carcases and animal products pose a negligible transmission risk. No special
disposal precautions are warranted.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999a, Import risk analysis: Importation of
dairy products for human consumption, AQIS, Canberra.
AQIS (Australian Quarantine and Inspection Service) 1999b, Importation of sausage casings into
Australia, import risk analysis, December 1999, AQIS, Canberra.
Hudson, J.R. 1968, Aust Vet J, vol. 44, pp. 123-129, cited in Windsor and Masiga 1977.
Hyslop, N. St G. 1959, Proc 16th Int vet Congress, pp. 535-538, cited in Windsor and Masiga
1977.
Laak, E.A. 1992, ‘Contagious bovine pleuropneumonia, a review’, Vet Quart, vol. 15, pp.104-110.
Nicholas, R., Bashiruddin, J., Ayling, R., & Miles, R. 2000, ‘Contagious bovine
pleuropneumonia: a review of recent developments’, Vet Bull, vol. 70, pp. 827-838.
Provost, A., Perreau, P., Breard, A., Le Goff, C., Martel, J.L., & Cottew, G.S. 1987, ‘Contagious
bovine pleuropneumonia’, Rev sci tech Off int Epiz, vol. 6, pp. 625-679, cited in Schneider et al
1994.
Schneider, H.P., van der Lugt, J.J., & Hubschle, O.J.B. 1994, ‘Contagious bovine
pleuropneumonia’, in Infectious diseases of livestock, Vol II, eds. Coetzer, J.A.W., Thomson,
G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp. 1485-1494.
Windsor, R.S., & Masiga, W.N. 1977, ‘Investigations into the epidemiology of contagious bovine
pleuropneumonia: the persistence of Mycoplasma mycoides sub-species mycoides in placenta,
urine and hay’, Bull An Health Prod Africa, vol. 25, pp. 357-370.
Experts contacted:
Dr Robin Nicholas
Veterinary Laboratories Agency (Weybridge)
New Haw, Addlestone
Surrey KT15 3NB
UK
P +44 1932 357379
F +44 1932 357423
[email protected]
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Epizootic lymphangitis
Agent:
Histoplasma capsulatum var farciminosum
Agent type:
Fungus
Persistence and inactivation:
Little information of relevance is available for this agent.
Bardelli and Ademollo (1927) reported virulence of the organism after desiccation in the
laboratory for 25 months. While Geering et al (1995) state that the organism persists for up to 15
days in the environment, Gabal and Hennager (1983) report much longer survival. Their study of
five recently isolated strains showed that temperature and moisture content of soil affected
viability. Maximum survival times in non-sterile soil and water for H. capsulatum were:
Temp (oC)
Soil (wks)
Water (wks)
-15
5
26
37
18
16
8
6
18
16
10
6
H. capsulatum is transmitted by direct contact, or via contamination of wounds by flies or
fomites. There are also reports of transmission from stallions to mares during copulation (Al-Ani
1999).
Al-Ani (1999) recommends slaughter of infected animals in the case of exotic outbreaks. No
references were found on the persistence of H. capsulatum in carcases.
Conclusions:
•
H.capsulatum is a persistent organism which can survive in the environment for
several months.
•
Given the persistence of H. capsulatum and potential for spread by flies and fomites,
burial or burning of infected carcases and contaminated animal products appears
warranted.
References:
Al-Ani, F.K. 1999, ‘Epizootic lymphangitis in horses: a review of the literature’, Rev sci tech Off
int Epiz, vol 18, pp. 691-699.
Bardelli, P., & Ademollo, A. 1927, ‘Sulla resistenza delle conture di Cryptococcus farciminosus.
Rivolta agli agenti fisici e chimici’, Annual Igiene, vol. 37, pp. 81-85, cited in Al-Ani 1999.
Gabal, M.A., & Hennager, S. 1983, ‘Study on the survival of Histoplasma farciminosum in the
environment’, Mykosen, vol. 26, pp. 481-484, 487.
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Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
Experts contacted:
Dr Falah Al-Ani
(formerly Jordan University of Science and Technology)
PO Box 620026
Irbid 21162
JORDAN
P +962 2 725 2985
F +962 2 727 0972
[email protected]
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Protozoa
Surra
Agent:
Trypanosoma evansi
Agent type:
Protozoan
Persistence and inactivation:
General characteristics: Trypanosoma evansi is a stercorarian trypanosome, i.e. there is no
intermediate host in which the organism multiplies and undergoes morphological transformation.
T. evansi is essentially transferred by mechanical means between mammalian hosts by biting
insects, notably Tabanus spp (horseflies) and Stomoxys spp (stable flies) (Connor 1994).
According to Brun (pers comm) there are few or no reports on the in vitro or in carcase survival
or inactivation of T. evansi in the literature. T. evansi is a fragile organism that disappears
quickly in the environment or after the death of the host (Geering et al 1995, Brun pers comm).
Several mainly foreign language papers on preservation for experimental purposes were
uncovered during this review but were considered of little relevance.
Hashemi Fesharki (1981) reported that infected blood kept at -70oC for eight years killed all of
five rats when inoculated at 1ml/rat.
Carcases and meat products: It is believed that T. evansi can be transmitted by ingestion of fresh
carcases. This has been confirmed in one case of a cat eaten by a canid, and the presence of
infection in hyenas and other scavengers strongly suggests transmission by this route. The chance
of human infection occurring via infected meat is considered negligible (Brun, pers comm).
The only objective study on survival in the carcase uncovered during this review was conducted
by Sarmah (1998), who examined the viability (by microscopic examination and mouse
inoculation) of T. evansi in the carcases of parasitaemic rats stored at room temperature post
mortem. All organisms were still viable at 4 hours; <1% were viable at 10 hours (although mice
were infected); and by 12 hours all parasites had degenerated.
Brun (pers comm) agrees that once the host is dead, conditions are rapidly untenable for the
parasite, and that the chance of survival in a carcase beyond 2-3 days is nil. The usual, ‘fail-safe’
process for disposing of experimental rodent carcases infected with T. evansi at the Swiss
Tropical Institute is to freeze/thaw and then incinerate them, although there is no empirical
evidence for the efficacy of this approach.
Milk and milk products: T. evansi may be directly transmitted through milk (Wang 1988).
Skins, hides and fibres: Infectivity of trypanosomes via skins, hides or fibres is extremely
unlikely given the fragile nature of the organism. In a draft assessment, AFFA (2001) has
assessed these products as posing no quarantine hazard for the importation of trypanosomiases.
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Semen/embryos: Trypanosoma equiperdum is found in the semen of horses (Brun et al 1998).
Wang (1998) reported that T. evansi could be directly transmitted through coitus.
Faeces: No references were found to the presence of T. evansi in faeces.
Conclusions:
•
T. evansi is a fragile organism which, under normal conditions, is unlikely to survive
away from an animal host for more than three days.
•
Spread between mammalian hosts mainly occurs with biting insects. However,
transmission can also spread with ingestion of an infected fresh carcase.
•
Fresh carcases should be disposed of so as to preclude scavenging.
•
Meat and other animal products pose a negligible transmission risk. No special
disposal precautions are warranted.
•
Semen should not be used.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
Brun, R., Hecker, H., & Lun, Z-R. 1998, ‘Trypanosoma evansi and T. equiperdum: distribution,
biology, treatment, and phylogenetic relationship (a review)’, Vet Parasitol, vol. 79, pp. 95-107.
Connor, R.J. 1994, ‘African animal trypanosomiases’, in Infectious diseases of livestock, Vol I,
eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp.
167-205.
Hashemi Fesharki, R. 1981, ‘Survival of Trypansomoa evansi, Theileria mutans, Babesia
bigemina and Anaplasma marginale following long-term maintenance at -70oC’, Arch Inst Razi,
vol. 32, pp. 47-50.
Sarmah, P.C. (1998), ‘Observation on survival of Trypanosoma evansi after death of the host’, J
Vet Parasitol, vol. 12(1), pp. 62-63.
Wang, Z-L. 1988, ‘The similarities and differences of the characteristics between T. equiperdum
and T. evansi’, Bul Vet Col (PLA), vol. 8, pp. 300-303 (in Chinese, cited in Brun et al 1998).
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Experts contacted:
Dr Reto Brun
Associate Professor
Swiss Tropical Institute
Protozoology Lab
Socinstr. 57
PO Box
CH-4002 Basel
SWITZERLAND
P +41 61 284 8231
F +41 61 271 8654
[email protected]
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Dourine
Agent:
Trypanosoma equiperdum
Agent type:
Protozoan
Persistence and inactivation:
General characteristics: Like T. evansi, Trypanosoma equiperdum is a stercorarian trypanosome,
having no intermediate host in which the organism multiplies and undergoes morphological
transformation. However T. equiperdum is unique among pathogenic trypanosomes in that it
does not even require a vector for transmission between hosts. T. equiperdum has developed the
capacity to survive in the genital tract and is spread directly by venereal means (Connor 1994).
Foals may also become infected, possibly at birth through infected vaginal discharges, udder
lesions, or via infected milk (Schulz 1935).
According to Brun (pers comm) there are few or no reports on the in vitro or in carcase survival
or inactivation of T. equiperdum in the literature. T. equiperdum is a fragile organism that
disappears quickly in the environment or after the death of the host (Geering et al 1995, Brun pers
comm). Several mainly foreign language papers on preservation for experimental purposes were
uncovered during this review but were considered of little relevance.
Carcases and meat products: No references were found on the survival of T. equiperdum in
carcases or meat products. It seems reasonable to extrapolate from the study by Sarmah (1998)
on survival of T. evansi in rat carcases, in which no viable organisms were found after 12 hours.
Brun (pers comm) agrees that once the host is dead, conditions are rapidly untenable for the
parasite, and that the chance of survival in a carcase beyond 2-3 days is nil.
Milk and milk products: The infection of foals suggests that transmission by milk may be
possible (Schulz 1935).
Skins, hides and fibres: Infectivity of trypanosomes via skins, hides or fibres is extremely
unlikely given the fragile nature of the organism. In a draft assessment, AFFA (2001) has
assessed these products as posing no quarantine hazard for the importation of trypanosomiases.
Semen/embryos: Trypanosoma equiperdum is transmitted by coitus. The organism is found in
the seminal fluid and mucous membranes of the genitalia of horses (Brun et al 1998).
Faeces: No references were found to the presence of T. equiperdum in faeces.
Conclusions:
•
T. equiperdium is a fragile organism which, under normal conditions, is unlikely to
survive away from an animal host for more than three days.
•
Disease spread occurs mainly by venereal means. Foals may possibly become
infected from contact with vaginal discharges or ingestion of infected milk.
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•
Carcases and animal products pose a negligible transmission risk. No special
disposal precautions are warranted.
•
Semen should not be used.
References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
Barrowman, P., Stoltsz, W.H., van der Lugt, J.J., & Williamson, C.C. 1994, ‘Dourine’, in
Infectious diseases of livestock, Vol I, eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C.,
Oxford University Press, Cape Town, pp. 206-212.
Brun, R., Hecker, H., & Lun, Z-R. 1998, ‘Trypanosoma evansi and T. equiperdum: distribution,
biology, treatment, and phylogenetic relationship (a review)’, Vet Parasitol, vol. 79, pp. 95-107.
Connor, R.J. 1994, ‘African animal trypanosomiases’, in Infectious diseases of livestock, Vol I,
eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape Town, pp.
167-205.
Sarmah, P.C. (1998), ‘Observation on survival of Trypanosoma evansi after death of the host’, J
Vet Parasitol, vol. 12(1), pp. 62-63.
Schulz, K. 1935, ‘Dourine or slapsiekte’, J Sth Afr Vet Med Ass, vol. 6, pp. 4-15, cited in
Barrowman et al 1994.
Experts contacted:
Dr Reto Brun
Associate Professor
Swiss Tropical Institute
Protozoology Lab
Socinstr. 57
PO Box
CH-4002 Basel
SWITZERLAND
P +41 61 284 8231
F +41 61 271 8654
[email protected]
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East Coast fever
Agent:
Theileria parva parva
Agent type:
Protozoan
Persistence and inactivation:
The only natural means of transmission of Theileria parva parva is via the tick Rhipicephalus
appendiculatus. Unlike the stercorarian trypanosomes such as T. evansi, however, the tick is
critical in the life cycle, being the host for sexual reproductive phase of the organism. Artificial
transmission can be effected by injecting ground-up tick (or its salivary glands) containing
sporozoites into the host. It can also be achieved by inoculation with suspensions of schizonts,
from spleen, lymph nodes, blood, or culture, although this method is more effective when the
cells originate from the donor (Lawrence et al 1994, Wilde 1967).
Destocking infected pastures of cattle for 15-18 months has eradicated infection, as this period
exceeds the maximum lifespan if infected ticks (Lawrence et al 1994).
According to Morzaria (pers comm) there are few or no reports on the in vitro or in carcase
survival or inactivation of T. parva parva in the literature, and none were uncovered during this
review. T. parva parva is a fragile organism that disappears quickly in the environment or after
the death of the host (Morzaria pers comm). Wilde (1967) noted that spleen used to induce
experimental infection is less infective if taken immediately before or at death than if taken
earlier. Several papers on preservation for experimental purposes were found but were
considered of little relevance.
Conclusions:
•
T. parva parva is a fragile organism which, under normal conditions, is unlikely to
survive away from an animal host or tick for more than two days.
•
Ticks are required for disease spread.
•
Carcases and animal products pose a negligible transmission risk. No special
disposal precautions are warranted.
References:
Lawrence, J.A., de Vos, A.J., & Irvin, A.D. 1994, ‘East Coast fever’, in Infectious diseases of
livestock, Vol I, eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press,
Cape Town, pp. 309-325.
Wilde, J.K.H. 1967, ‘East Coast fever’, in Advances in Veterinary Science Volume II, eds
Brandly, C.A., & Cornelius, C., Academic Press, New York, pp. 207-259.
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Experts contacted:
Dr Subhash Morzaria
International Livestock Research Institute
PO Box 30709
Nairobi
KENYA
P +254 2 630743
F +254 2 631499
[email protected]
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Equine babesiosis (piroplasmosis)
Agent:
Babesia equi and Babesia caballi
Agent type:
Protozoa
Persistence and inactivation:
As with Theileria parva parva, an intermediate tick host is critical to the life cycles of Babesia
equi and B. caballi. Several ticks are implicated in transmission (Rhipicephalus spp, Hyalomma
spp, and Dermacentor spp). Disease has been induced by inoculation of infected blood into
susceptible animals, and iatrogenically via contaminated needles (de Waal and van Heerden
1994).
No reports of the in vitro or in carcase survival or inactivation of B. equi or B. caballi were found
in the literature. As with other protozoa, preservation of live organisms is difficult. Babesia
bovis can be detected in heart, lung and kidney up to eight hours after death, and up to 28 hours in
brain (Radostits et al 2000). Several papers on preservation for experimental purposes were
found but were considered of little relevance.
Conclusions:
•
Babesia equi and Babesia caballi are fragile organisms which, under normal
conditions, are unlikely to survive from an animal host or tick for more than two
days.
•
Ticks are required for disease spread.
•
Carcases and animal products pose a negligible transmission risk. No special
disposal precautions are warranted.
References:
De Waal, D.T., & van Heerden, J. 1994, ‘Equine babesiosis’, in Infectious diseases of livestock,
Vol I, eds. Coetzer, J.A.W., Thomson, G.R., & Tustin, R.C., Oxford University Press, Cape
Town, pp. 295-304.
Radostits, O.M., Gay, C.G., Blood, D.C., & Hinchcliff, K.W. (eds) 2000, Veterinary medicine,
ninth edition, W. B. Saunders Company Ltd, London.
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Multicellular parasites
Sheep scab
Agent:
Psoroptes ovis
Agent type:
Mite
Persistence and inactivation:
Psoroptes ovis is an obligate parasite. Its survival off the host has been the subject of widely
differing estimation and opinion (summarised in O’Brien et al 1994). The study coming closest
to mimicking natural conditions appears to be that of O’Brien et al (1994), in which the mites
retained infectivity for up to 16 days under ambient conditions in Ireland. Survival and
infectivity was consistent across all seasons and in the refrigerator. Some mite eggs hatched after
a week off the host when subsequently placed in an incubator. The eggs hatched in 1-3 days after
placement in the incubator or not at all.
Using artificial environments, other workers have reported longer survival times (for example, 48
days in the laboratory for a bovine strain by Liebisch et al 1985). Smith et al (1999) showed that
temperature and humidity affected survival of P. ovis. Maximum survival times of P. ovis,
maintained in chambers and supplied with distilled water, were 7-8 days at 24-26oC, and 15-18
days at 2-9oC. The closely related species P. caniculi from rabbits appeared to have significantly
lower survival below a relative humidity of 65-75% at 30oC.
P. ovis is susceptible to a number of ectoparasiticides, and is usually treated by topical treatment
using an organophosphate (e.g. diazinon 0.01%, propetamphos 0.0125%). Ivermectin may also
be effective (Radostits et al 2000).
P. ovis is an external parasite and therefore not associated with meat, milk, semen/embryos or
faeces. Survival of the mite on carcases might be expected to fall within the range of estimates
described above, but no specific studies on survival on carcases were found during this review
(the lack of data was confirmed by Wall, pers comm).
In a draft assessment, AFFA (2001) has concluded that the risk of spreading P. ovis by raw hides
or wool is slight. P.ovis is unlikely to survive the drying process of hides and scouring of wool.
Conclusions:
•
P. ovis is a sensitive mite which, under natural conditions, is unlikely to survive for
more than three weeks away from an animal host.
•
Close physical contact is required for transmission.
•
Processed skins, scoured wool and other animal products pose a negligible
transmission risk.
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References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
Liebisch, A., Olbrich, S., & Deppe, M. 1985, ‘Survival of P. ovis, P. cuniculi, C. bovis when
separated from the host animal’, Dtsch Tieraerztl Wochenschr, vol. 92, pp. 165-204, cited in
O’Brien et al 1994 and Smith et al 1999.
O’Brien, D.J., Gray, J.S., & O’Reilly, P.F. 1994, ‘Survival and retention of infectivity of the mite
Psoroptes ovis off the host’, Vet Res Comm, vol. 18, pp. 27-36.
Radostits, O.M., Gay, C.G., Blood, D.C., & Hinchcliff, K.W. (eds) 2000, Veterinary medicine,
ninth edition, W. B. Saunders Company Ltd, London.
Smith, K.E., Wall, R., Berriatua, E., & French, N.P. 1999, ‘The effects of temperature and
humidity on the off-host survival of Psoroptes ovis and Psoroptes cuniculi ’, Vet Parasitol, vol.
83, pp. 265-275.
Experts contacted:
Professor Richard Wall
School of Biological Sciences
University of Bristol
Bristol BS8 1UG
UK
P +44 117 928 9182
F +44 117 925 7374
[email protected]
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Tropilaelaps mite
Agent:
Tropilaelaps clareae
Agent type:
Mite
Persistence and inactivation:
AUSVETPLAN (DPIE 1996) notes that Tropilaelaps clareae persists only in a hive with adult
bees and live brood. Adult mites do not survive more than 2-4 days away from bee brood.
Gravid females survive only a short period on the adult bee, because their unspecialised
chelicerae cannot pierce the integument of the host. They die within two days unless they can
deposit their eggs (Sammataro et al 2000).
The literature contains numerous references to control of T. clareae populations using
chemotherapy. Control with chemotherapeutic agents is difficult, because many of the mites are
physically partitioned from the chemical, necessitating multiple treatments over several weeks to
combat the ongoing emergence of parasites from the brood (Burgett and Kitprasert 1990).
A number of treatments have been shown to reduce mite populations, including formic acid,
sulphur and chlorobenzilate (Sammataro et al 2000). A slow-release fluvalinate formulation
appears to be the preferred treatment (Burgett and Kitprasert 1990).
Conclusions:
•
AUSVETPLAN stipulates the destruction of infected hives by either burning or
treatment with an insecticide or solvent followed by burial.
•
Given the low unit value of bees and their products and the relative ease of disposal,
there is no obvious reason to consider lower cost, higher risk disposal methods.
References:
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Bee diseases, DPIE, Canberra.
Burgett, D.M., & Kitprasert, C. 1990, ‘Evaluation of Apistan as a control for Tropilaelaps clareae
(Acari: Laelapidae), an Asian honey bee brood mite parasite’, Am Bee J, vol. 130, pp. 51-53.
Sammataro, D., Gerson, U., & Needham, G. 2000, ‘Parasitic mites of honey bees: life history,
implications, and impact’, Annu Rev Entomol, vol. 45, pp. 519-548.
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Varroa mites
Agent:
Varroa destructor and Varroa jaobsoni
Agent type:
Mite
Persistence and inactivation:
Varroa destructor has only recently been described as a separate species from Varroa jacobsoni
after Anderson and Trueman (2000) demonstrated significant genetic variation. Varroa jacobsoni
reproduces only on Apis cerana and is thus not a serious problem for managed honey bee
colonies (Apis mellifera). Two haplotypes of V. destructor cause significant damage to A.
mellifera (Anderson and Trueman 2000). The literature prior to this time therefore contains a
mixture of descriptions of both species under the name V. jacobsoni (Anderson pers comm).
Varroa mites are transmitted when beekeepers transfer colonies or brood between colonies, or
when bees rob each other or meet at flowers. The feral bee population may act as a reservoir of
infection (Anderson pers comm).
As with the other mites of bees considered in this review, Varroa spp are obligate ectoparasites
(Anderson and Trueman 2000). Varroa can survive off the host for 18-70 hours (Sammataro et al
2000). Anderson (1997) found that adult female V. jacobsoni did not survive longer than 9 days
in leafcutter bee nest material at 5oC. AUSVETPLAN states that mites will die in three days
without food (DPIE 1996). Anderson (1994), examining the survival of adult female mites in the
presence or absence of A. cerana and A. mellifera worker brood, reported survival of mites away
from pupae for up to 96 hours.
A slow-release fluvalinate formulation appears to be the preferred treatment for Varroa spp,
although resistance to this chemical has been recorded since 1995 in Italy. Other treatments
include coumaphos, formic acid, oxalic and lactic acids (Sammataro et al 2000). The
effectiveness of a gel formulation of thymol has recently been demonstrated (Mattila and Otis
1999).
Destruction of bees, combs and hive components as well as feral bee colonies where present is the
only method for eradication, and would need to be instituted very quickly in case of an incursion.
Conclusions:
•
AUSVETPLAN stipulates the destruction of infected hives by either burning or
treatment with an insecticide or solvent followed by burial.
•
Given the low unit value of bees and their products and the relative ease of disposal,
there is no obvious reason to consider lower cost, higher risk disposal methods.
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References:
Anderson, D.L. 1994, ‘Non-reproduction of Varroa jacobsoni in Apis mellifera colonies in Papua
New Guinea and Indonesia’, Apidologie, vol. 25, pp. 412-421.
Anderson, D.L. 1997, Evaluation of quarantine risks associated with importations of the
Leafcutter Bee (Megachile rotundata) to Australia from Canada. Report on a consultancy
conducted in Canada and the United States from 30 June to 15 July 1997. CSIRO Division of
Entomology, Canberra.
Anderson, D.L., & Trueman, J.W.H. 2000, ‘Varroa jacobsoni (Acari: Varroidae) is more than
one species’, Exp Appl Acarol, vol. 24, pp. 165-189.
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Bee diseases, DPIE, Canberra.
Mattila, H.R., & Otis, G.W. 1999, ‘Trials of Apiguard, a thymol-based miticide. Part 1. Efficacy
for control of parasitic mites and residues in honey’, Am Bee J, vol. 139, pp. 947-952.
Sammataro, D., Gerson, U., & Needham, G. 2000, ‘Parasitic mites of honey bees: life history,
implications, and impact’, Annu Rev Entomol, vol. 45, pp. 519-548.
Experts contacted:
Dr Denis Anderson
CSIRO Entomology
GPO Box 1700
Canberra ACT 2600
AUSTRALIA
P 02 6246 4148
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Acarapis (tracheal) mite
Agent:
Acarapis woodi
Agent type:
Mite
Persistence and inactivation:
Acarapis woodi is an obligate parasite that primarily infests the tracheae and air sacs of adult
honey bees (Apis mellifera). Mites disperse by questing on bee setae when the host bee is 15-25
days old. Survival of mites may not exceed a few hours during this period off the host, as mites
are very sensitive to desiccation and starvation. Ambient temperature, humidity and state of
nourishment are critical factors in the survival time (Sammataro et al 2000).
A. woodi poses problems for chemical control, because suitable treatments must reach bee
tracheae via a volatile compound, be inhaled by the bee, and be lethal only to the mite. Residues
in bee products must also be avoided. The only product registered in the United States is pure
menthol crystals. Amitraz and formic acid are also used as treatments elsewhere (Sammataro et
al 2000). Flumethrin, fluvalinate, and methyl palmitate can be used to reduce mite populations.
Residues in honey and beeswax have been identified following chemotherapy.
Conclusions:
•
AUSVETPLAN stipulates the destruction of infected hives by either burning or
treatment with an insecticide or solvent followed by burial.
•
Given the low unit value of bees and their products and the relative ease of disposal,
there is no obvious reason to consider lower cost, higher risk disposal methods.
References:
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Bee diseases, DPIE, Canberra.
Sammataro, D., Gerson, U., & Needham, G. 2000, ‘Parasitic mites of honey bees: life history,
implications, and impact’, Annu Rev Entomol, vol. 45, pp. 519-548.
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Screw-worm fly
Agent:
Chrysomya bezziana, Cochliomyia hominivorax
Agent type:
Fly
Persistence and inactivation:
Screw-worm flies lay eggs on the dry edge of wounds or body orifices. Eggs hatch within 12-20
hours, and the first instar larvae burrow into the wound and begin to feed. Moults to second and
third instar larval stages occur after successive periods of 24 hours. The larvae feed until 6-7
days old then drop off to pupate in the soil. Pupation lasts for a week to two months, depending
on ambient conditions. Adult males and female flies emerge and commence mating, with females
laying the first batch of eggs after 6-7 days. The life cycle can be completed in 20 days in
optimal conditions. The average adult lifespan of the SWF is 21 days (Spradbery 1994, Geering
et al 1995).
Screw-worm flies will probably not survive in open areas subject to dry heat, unless vegetation is
available for shade and carbohydrate. SWF prefers an ambient temperature of 20-30oC, will not
move at <10oC and may not mate at 10-16oC. It does not survive in areas that experience frosts
(Geering et al 1995).
The preferred prophylactic treatment for livestock in the face of a screw-worm fly incursion is
ivermectin. A recent study showed that ivermectin boluses protected cattle from 14 to 21 days
after treatment and inhibited fly breeding in dung, although with negative impacts on dung beetles
(Wardaugh et al 2001).
Larval screw-worm flies are unlikely to persist long on carcases or skins, as they rely on blood for
nourishment. AFFA (2001) determined that there is some quarantine risk posed by unprocessed
hides from SWF-endemic countries. It noted that treatment with methyl bromide, ethylene oxide
or other insecticidal gases would be effective with individual hides but was unlikely to penetrate
layers of hides, so post-border processing in quarantine would be necessary.
AUSVETPLAN (DPIE 1996) notes that the only animal product necessary to treat is faeces, as it
may contain pupating larvae. The preferred product is bromophos plus chlorfenvinphos, with
phosmet, crotoxyphos or cypermethirn plus chlorfenvinphos as alternatives.
Conclusions:
•
Screw-worm fly is an insect pest that can breed on a range of animal hosts.
•
Australia’s response plan for a screw worm fly incursion does not include the
slaughter of livestock or destruction of animal products.
•
To prevent transportation of the fly to new regions, and to minimise breeding
grounds for the fly, disinsection of faeces on plant and equipment may be necessary.
Unprocessed hides being exported from a control area may also require treatment.
•
Meat and milk pose a negligible transmission risk. No special disposal precautions
are warranted.
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References:
AFFA (Department of Agriculture Fisheries and Forestry – Australia) 2001, Import risk analysis.
Skins and hides. Draft report, August 2001, AFFA, Canberra.
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Screw-worm fly, DPIE, Canberra.
Geering, W.A., Forman, A.J., & Nunn, M.J. 1995, Exotic diseases of animals: a field guide for
Australian veterinarians, Australian Government Publishing Service, Canberra.
Spradbery, J.P. 1994, ‘Screw-worm fly: a tale of two species’, in Agricultural Zoology Reviews,
vol. 6, October 1994, ed. Evans, K., Intercept Ltd, Andover, pp. 1-61.
Wardaugh, K.G., Mahon, R.J., & Ahmad, H.B. 2001, ‘Efficacy of macrocyclic lactones for the
control of larvae of the Old World Screw-worm Fly (Chrysomya bezziana)’, Aust vet J, vol. 79,
pp. 120-124.
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Braula fly
Agent:
Braula coeca
Agent type:
Fly
Persistence and inactivation:
Braula coeca is a dipterous fly and commensal of honey bees. Larvae pupate in tunnels below
cappings, and on emergence, the adult flies attach to adult worker bees, queen bees and
sometimes drones. DPIE (1996) states that B. coeca is not known to survive in the absence of
adult bees, although it will survive in a colony without brood. No references to persistence off
the host were found during this review.
B. coeca is spread by swarming or drifting colonies and by the transfer of colonies or hive
components (DPIE 1996).
Kulincevic et al (1991) reported that fluvalinate as an aerosol or fumigant killed significant
numbers of Braula but that amitraz was not effective.
Conclusions:
•
AUSVETPLAN stipulates the destruction of infected hives by either burning or
treatment with an insecticide or solvent followed by burial.
•
Given the low unit value of bees and their products and the relative ease of disposal,
there is no obvious reason to consider lower cost, higher risk disposal methods.
References:
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Bee diseases, DPIE, Canberra.
Kulincevic, J.M., Rinderer, T.E., & Mladjan, V.J. 1991, ‘Effects of fluvalinate and amitraz on bee
lice (Braula coeca Nitzsch) in honey bee (Apis mellifera L) colonies in Yugoslavia’, Apidologie,
vol. 22, pp. 43-47.
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Small hive beetle
Agent:
Aethina tumida
Agent type:
Beetle
Persistence and inactivation:
Limited research exists on small hive beetle, as it has only been described as a pathogen of honey
bee colonies in the Western Hemisphere since 1998. Larvae of A. tumida migrate from combs to
pupate in surrounding soil. Studies in the US found beetle larvae, pupae and adults at a depth of
1-20cm in soil around hive entrances. Eighty percent of the beetles were within 30cm, and none
were further than 180cm from the hive (Pettis and Shimanuki 2000). Development from egg to
adult takes 38-81 days (Lundie 1940). Adults survived for up to five days without food or water
in moderate summer temperatures (Pettis and Shimanuki 2000), but could survive much longer if
provided food (e.g. greater than 60 days reported by Lundie (1940) for beetles fed honey and
pollen). Elzen et al (1999) observed small hive beetle mortalities as high as 90.2% using 10%
coumaphos in plastic strips.
Conclusions:
•
Small hive beetle would be more difficult to eradicate than other bee pests because
of its capacity to survive free of the colony. However, the destruction of both hives
and feral bee colonies would be an eradication option for Australia in the case of an
isolated incursion.
•
The preferred methods for destroying infected hives are either burning or treatment
with an insecticide or solvent followed by burial.
•
Given the low unit value of bees and their products and the relative ease of disposal,
there is no obvious reason to consider lower cost, higher risk disposal methods.
References:
DPIE (Department of Primary Industries and Energy) 1996, Australian Veterinary Emergency
Plan, AUSVETPLAN Edition 2.0, Disease Strategy Bee diseases, DPIE, Canberra.
Elzen, P.J., Baxter J.R., Westervelt, D., Randall, C., Delaplane, K.S., Cutts, L., & Wilson, W.T.
1999, ‘Field control and biology studies of a new pest species, Aethina tumida Murray
(Coleoptera, Nitidulidae), attacking European honey bees in the Western hemisphere’,
Apidologie, vol. 30, pp. 361-366.
Lundie, A.E. 1940, The small hive beetle, Aethina tumida, Sci Bull 220, Department of
Agriculture and Forestry, Union of South Africa, 30pp, cited in Elzen et al 1999.
Pettis, J.S., & Shimanuki, H. 2000, ‘Observations on the small hive beetle, Aethina tumida
Murray, in the United States’, Am Bee J, vol. 140, pp 152-155.
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Experts contacted:
Bruce White
NSW Agriculture
Locked Bag 11
WINDSOR NSW 2756
P 02 4577 0600
[email protected]
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Trichinellosis
Agent:
Trichinella spiralis
Agent type:
Nematode
Persistence and inactivation:
General characteristics: T. spiralis is a nematode. Adults are present in the intestine. Fertilised
females burrow into the villi and produce L1 larvae, which enter the lymphatics and travel via the
bloodstream to the skeletal muscles. They penetrate muscle cells and become encapsulated by the
host. The larvae resume development when the muscle tissue is eaten by another host. Larvae
remain infective in living hosts for many years. Transmission also occurs by ingestion of fresh
faeces from animals with a patent infection.
Trichinella have a wide range of hosts, including some aquatic mammals. Crustaceans and fish
may have a role as transport hosts in these infections (Urquhart et al 1987).
Carcases and meat products: Trichinella is spread by feeding of swill containing under-cooked
pig flesh to other pigs. Rodents and other rodents also act as a reservoir of infection.
According to Urquhart et al (1987), Trichinella maintain infectivity in decomposing carcases for
several months. Jovic et al (2000) studied the survival of T. spiralis larvae in 700g pieces of pig
muscle buried at depths of 30, 50 and 100cm at a temperature between 4-13oC. Infectivity was
maintained throughout the 91-day experimental period at all depths. Typical putrefaction with
marked odour and tissue decay was not observed, reflecting the low temperatures in the soil. The
authors compared their results with those of Modic (1976), who reported survival of larvae
between 25 and 40 days at 30oC.
The International Commission on Trichinellosis has published standards for the control of
Trichinella in pork for human consumption (Gamble et al 2000). The ICT states that there are
three acceptable methods for inactivating Trichinella spiralis in meat: cooking, freezing, and
irradiation.
For cooking, the ICT recommends heating according to one of a series of time and temperature
combinations set out by the United States Department of Agriculture’s Code of Federal
Regulations (USDA, 1990) (see table).
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Min. int. temp. (oC)
Min. time
49.0
50.0
51.1
52.2
53.4
54.5
55.6
56.7
57.8
58.9
60.0
61.1
62.2
21h
9.5h
4.5h
2h
1h
30min
15min
6min
3min
2min
1min
1min
instant
The standards further stipulate that the time to raise the product from 15.6oC to 49.0oC should not
exceed 2h unless the product is cured or fermented, and that steps be taken to ensure all parts of
the product are heated through. In the absence of proper temperature and time control and
monitoring systems, the meat should be checked to ensure colour has changed from pink to grey
throughout, and the texture such that muscle fibres are easily separated.
Smoking, drying or curing pork does not necessarily inactivate Trichinella larvae (Urquhart et al
1987).
For freezing, the ICT again recommends guidelines from the USDA’s Code of Federal
Regulations (USDA 1990). The Code specifies the following minimum times at various
maximum temperatures:
Max. internal temp. (oC)
Group 1* (days)
Group 2** (days)
-15
-23
-29
20
10
6
30
20
12
* pieces not greater than 15cm thickness
** pieces between 15cm and 69cm thickness
Where proper time and temperature control and monitoring systems are not available, pieces of
meat up to 15cm thick should be frozen solid (>15oC) for at least three weeks, and for meat 1569cm thick, for at least four weeks. These recommendations apply to pig and horse meat, but not
game meats, as these often harbour freeze-resistant species of Trichinella. (Pigs in certain
regions may also be infected with freeze-resistant Trichinella.)
A broader range of time/temperature options for freezing is provided by Kotula et al (1990), who
studied the inactivation of T. spiralis in pork at temperatures ranging from -193oC to -1oC, for
periods from 1 second to 182 days. Thermal death curves included the following points:
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Temp
o
C
Predicted thermal death
time
Predicted 99% confidence
limit
Predicted 99.999% confidence
limit
-10
-15
-20
4.0 days
64 min
8 min
266 days
63 hrs
48 min
NR
180 days
21 days
NR not reported
For irradiation, the ICT recommends 0.3 kGy (for use on sealed packaged food only).
Milk and milk products: No reports were found of Trichinella larvae in milk or milk products.
Skins, hides and fibres: These products are highly unlikely to pose an infective risk.
Semen/embryos: No reports were found of Trichinella larvae in semen or on embryos.
Faeces: Larvae may be found in faeces of patent, infected animals (Urquhart et al 1987). No
references were found regarding persistence of larvae in faeces.
Conclusions:
•
T. spiralis larvae encapsulated in muscle cells remain viable well beyond the life of
the animal host.
•
Transmission occurs with the ingestion of muscle tissue containing T. spiralis cysts
or the ingestion of fresh faeces from an infected animal.
•
Fresh carcases must be disposed of in a manner that precludes scavenging.
•
Infected meat poses a disease transmission and human health risk. T. spiralis cysts
can be destroyed by freezing, heating and/or irradiation, but this must be done
under controlled conditions to ensure inactivation of the parasite.
•
Skins and milk pose a negligible transmission risk. No special disposal precautions
are warranted.
References:
Gamble, H.R., Bessonov, A.S., Cuperlovic, K., Gajadhar, A.A., van Knapen, F., Noeckler, K.,
Schenone, H., & Zhu, X. 2000, ‘International Commission on Trichinellosis: recommendations
on methods for the control of Trichinella in domestic and wild animals intended for human
consumption’, Vet Parasitol, vol. 93, pp. 393-408.
Jovic, S., Djordjevic, M., Kulisic, Z., Pavlovic, S., & Radenkovic, B. 2001, ‘Infectivity of
Trichinella spiralis larvae in pork buried in the ground’, Parasite, vol. 8, pp. S213-S215.
Kotula, A.W., Sharar, A.K., Paroczay, E., Gamble, H.R., Murrell, K.D., & Douglass, L. 1990,
‘Infectivity of Trichinella spiralis from frozen pork’, J Food Protection, vol. 53, pp. 571573,626.
Urquhart, G.M., Armour, J., Duncan, J.L., Dunn, A.M., & Jennings, F.W. 1987, Veterinary
parasitology, Longman Scientific and Technical, Avon, UK, 286 pp.
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USDA (United States Department of Agriculture) 1990, Code of Federal Regulations, Animals
and animal products, Office of the Federal Registrar, Government Printing Office, Washington
D.C., vol. 9, pp. 212-230, cited in Gamble et al 2000.
Experts contacted:
Dr Alvin Gajadhar
Director
Centre for Animal Parasitology
Canadian Food Inspection Agency Laboratory
116 Veterinary Rd
Saskatoon, Sask. S7N 2R3
CANADA
P +1 306 975 5344
F +1 306 975 5711
[email protected]
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Appendix 1
Example search strategy
DATABASES SEARCHED
Database: Medline
Version: PubMed ((http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed)
Files searched: all
Date searched: 14-Sep-01
Search strategy:
babesiosis [MESH] AND horses [MESH]
equine babesiosis OR equine piroplasmosis
1 OR 2
survival OR persist* OR inactivat* OR disinfect*
3 AND 4
review OR review literature OR review of reported cases OR review, academic OR review,
multicase OR review, tutorial OR scientific integrity review [PUBLICATION TYPE]
3 AND 6
5 OR 7
Database: CAB Abstracts
Version: SilverPlatter on WebSpirs
Files searched: 1972-2001/07
Date searched: 14-Sep-01
Search stategy:
equine babesiosis OR equine piroplasmosis
babesia equi OR babesia caballi
2 OR 3
(virus OR agent OR disease) NEAR (survival OR persist* OR destruct*)
(viral OR virus OR agent)NEAR (inactivat* OR disinfect* OR persist*)
4 OR 5
3 AND 6
review in ti,ab
3 AND 8
7 OR 9
Database: Agricola
Version: SilverPlatter on WebSpirs
Files searched: 1992-2001/06
Date searched: 14-Sep-01
Search strategy: As above for CAB Abstracts
Database: ABOA
Version: SilverPlatter on WebSpirs
Files searched: 1975Date searched: 14-Sep-01
Search strategy: As above for CAB Abstracts
Persistence of Disease Agents
Revised December 2003
170
Scott Williams Consulting Pty Ltd
Database: Current Contents
Version: OVID at http://gateway.ovid.com/
Files searched: AGR, LIFE Week 29 2000 - week 38 2001
Date searched:14-Sep-01
Search strategy: As above for CAB Abstracts
Persistence of Disease Agents
Revised December 2003
171
Scott Williams Consulting Pty Ltd